Membrane-spanning nanopores

ABSTRACT

A membrane-spanning nanopore is provided that comprises:
         i. at least one scaffold polynucleotide strand;   ii. a plurality of staple polynucleotide strands; and   iii. at least one hydrophobically-modified polynucleotide strand, wherein the at least one hydrophobically-modified polynucleotide strand comprises a polynucleotide strand and a hydrophobic moiety; wherein each of the plurality of staple polynucleotide strands hybridises to the at least one scaffold polynucleotide strand to form the three-dimensional structure of the membrane-spanning nanopore, and wherein the at least one hydrophobically-modified polynucleotide strand hybridises to a portion of the at least one scaffold polynucleotide strand, the membrane-spanning nanopore defining a central channel with a minimum internal width of at least about 5 nm.

CROSS-REFERENCE TO RELATED APPLICATIONS

This application is a divisional application under 35 U.S.C. § 121 of co-pending U.S. application Ser. No. 16/317,085 filed Jan. 11, 2019, which is a 35 U.S.C. § 371 National Phase Entry Application of International Application No. PCT/GB2017/052089 filed Jul. 14, 2017, which designates the U.S. and claims benefit of foreign priority under 35 U.S.C. § 119(a) of GB provisional application No. 1612458.8 filed Jul. 14, 2016, the contents of which are incorporated herein by reference in their entireties.

SEQUENCE LISTING

The instant application contains a Sequence Listing which has been submitted electronically in XML format and is hereby incorporated by reference in its entirety. Said XML copy, created on Sep. 6, 2022, is named P42206USA_seqlist.xml and is 316,260 bytes in size.

TECHNICAL FIELD

The present invention relates to novel membrane nanostructures and their uses. In particular it relates to wide-channel membrane nanopores in the applications of protein sensing and molecular gate creation.

BACKGROUND

Nanopores are membrane spanning polymers and complexes that form a channel in a membrane through which ions and certain molecules may pass. The minimum diameter of the channel is typically in the nanometre (10⁻⁹ metre) range hence giving certain of these polypeptides the name ‘nanopores’.

Nanopores bound in membranes have many potential uses. One example is the use of nanopores as sensors to analyse biomolecules in a label-free and portable fashion. In an embodiment of this approach, an electrical potential is applied across a membrane-bound nanopore causing ions to flow through the channel. This flow of ions can be measured as an electrical current. Suitable electrical measurement techniques using single channel recording have been described, for example, in WO 2000/28312 and D. Stoddart et al., Proc. Natl. Acad. Sci., 2010, 106, 7702-7. Multi-channel recording techniques have also been described, for example, in WO 2009/077734 and International Application WO-2011/067559. Optical measurements may be combined with electrical measurements (Soni G V et al., Rev Sci Instrum. 2010 January; 81(1):014301). Alternatively flow of ions through the membrane-bound nanopore may be achieved by providing an ionic gradient across the membrane.

Other potential uses are in the provision of functional interconnected networks of droplets joined by droplet interface bilayers containing nanopores that exploit a variety of membrane pumps, channels and pores to act as light sensors, batteries, and electrical devices (see for example Holden, M, A, et al, J. Am, Chem, Soc. 129, 8650-8655 (2007); 25 Maglia, G. et al. Nat. Nanotechnol, 4, 437-440 (2009)). Another use is the provision of droplet encapsulates or multisomes for use as drug delivery vehicles, such as disclosed by International Application PCT/GB2012/052736.

Individual molecules passing, binding or lodging inside a nanopore reduces the ion flow through the channel to yield an ion current read-out (see, for example, Liu X., Mihovilovic Skanata M., Stein D. Nat. Commun. 6, 6222 (2015); Lindsay S. Nat. Nanotechnol. 11, 109-111 (2016); Howorka S., Siwy Z. Chem. Soc. Rev. 38, 2360-2384 (2009); Wang Y., Zheng D., Tan Q., Wang M. X., Gu L. Q. Nat. Nanotechnol. 6, 668-674 (2011); and Wei R. S., Gatterdam V., Wieneke R., Tampe R., Rant U. Nat. Nanotechnol. 7, 257-263 (2012)). The degree of reduction in ion flow, as measured by the reduction in electrical current, is indicative of the size of the obstruction within, or in the vicinity of, the pore. The measured electrical current can therefore be used as a measure of the size or degree of obstruction to the channel. The changes in electrical current can be used to identify that a molecule, or part of a molecule, has bound at or near the pore (molecular sensing), or in certain systems, it can be used to determine the identity of a molecule that is present within the pore based on its size. This principle is used in nanopore based nucleic acid sequencing.

Adapting the sensing approach to specific analytes relies on nanopores of tailored structure. Prominent examples are the membrane pores MspA and α-hemolysin which have a protein structure and whose narrow lumen matches the size of DNA nucleotides and thus allow electrical sequencing of individual translocating DNA strands (Cherf G. M., Lieberman K. R., Rashid H., Lam C. E., Karplus K., Akeson M. Nat. Biotechnol. 30, 344-348 (2012); Manrao E. A., et al. Nat. Biotechnol. 30, 349-353 (2012); and Quick J., et al. Nature 530, 228-232 (2016)), or nucleotides (Clarke J., Wu H. C., Jayasinghe L., Patel A., Reid S., Bayley H. Nat. Nanotechnol. 4, 265-270 (2009)).

Expanding the powerful sensing principle to protein analytes of diagnostic or environmental importance that remain in their native folded state requires a different set of nanopores.

To be useful as a sensor for folded proteins or other large biomolecules, suitable membrane channels formed by a nanopore should meet certain criteria, namely:

-   -   1) the channel lumen should be at least about 5 nm wide to         accommodate the large biomolecule molecules inside the channel         lumen; binding of the large biomolecule within the channel         results in higher read-out sensitivity than when analytes bind         at the pore entrance;     -   2) the pores should be structurally defined to attain a constant         base level in the electrical read-out (i.e. reduce background         noise); and     -   3) the pore dimensions should be easily tunable to adapt them to         different biomolecule sizes.

To date none of the existing biological or engineered pores fulfill all of these criteria.

In terms of potential protein pores, the narrow α-hemolysin pore can be tailored to achieve binding of selected proteins at the pore entrance (Movileanu L., Howorka S., Braha O., Bayley H. Nat. Biotechnol. 18, 1091-1095 (2000); Rotem D., Jayasinghe L., Salichou M., Bayley H. J. Am. Chem. Soc. 134, 2781-2787 (2012)) but this approach is not generic. Furthermore, perfringolysin and related pores have a diameter of at least 20 nm yet their size is heterogeneous (Dang T. X., Hotze E. M., Rouiller I., Tweten R. K., Wilson-Kubalek E. M. J. Struct Biol. 150, 100-108 (2005)). By comparison, ClyA has a defined width of 4.5 nm; its diameter is not easily tunable (Maglia G., Henricus M., Wyss R., Li Q., Cheley S., Bayley H. Nano Lett. 9, 3831-3836 (2009); Soskine M., Biesemans A., Maglia G. J. Am. Chem. Soc. 137, 5793-5797 (2015). Engineering protein pores de novo (Dang et al, ibid.) is not an option due to the difficulty of predicting the final outcome of folding polypeptides.

Sufficiently wide nanopores can also be fabricated from inorganic materials (see, for example, Wei R. S., Gatterdam V., Wieneke R., Tampe R., Rant U. Nat. Nanotechnol. 7, 257-263 (2012); Dekker C. Nat. Nanotechnol. 2, 209-215 (2007); Miles B. N., Ivanov A. P., Wilson K. A., Dogan F., Japrung D., Edel J. B. Chem. Soc. Rev. 42, 15-28 (2013); and Yusko E. C., et al. Nat. Nanotechnol. 6, 253-260 (2011)) but they are not compatible with hydrophobic membrane format that is standard for portable analytical nanopore devices (Quick J., et al. Nature 530, 228-232 (2016)).

Pores composed of nucleic acid duplexes, in particular DNA duplexes, represent another possibility for sensing nanopores. DNA nanopores have recently been obtained from a structural core of six hexagonally arranged, interlinked DNA duplexes that enclose a hollow channel (see, for example, Douglas S. M., Marblestone A. H., Teerapittayanon S., Vazquez A., Church G. M., Shih W. M. Nucleic Acids Res. 37, 5001-5006 (2009); Zheng J., et al. Nature 461, 74-77 (2009); Rothemund P. W. Nature 440, 297-302 (2006); Fu J., et al. Nat. Nanotechnol. 9, 531-536 (2014); Burns J. R., et al. Angew. Chem. Int. Ed. 52, 12069-12072 (2013); and Seifert A., Göpfrich K., Burns J. R., Fertig N., Keyser U. F., Howorka S. ACS Nano 9, 1117-1126 (2015)). Membrane insertion was achieved through equipping the pores exterior with hydrophobic lipid anchors. However, these pores have a narrow lumen of no more than 2 nm in diameter. Another drawback was structural instability leading to an 80% channel closure at standard experimental conditions of high transmembrane voltages (Zheng et al, ibid.).

The modular construction principle for DNA nanopores has enabled customized pore diameter (Göpfrich et al, Nano. Lett., 15(5), 3134-3138 (2015); WO 2013/083983) and installation of a controllable gate to regulate transport (Burns J. R., Seifert A., Fertig N., Howorka S. A., Nat. Nanotechnol. 11, 152-156 (2016)).

A general challenge remains to insert the negatively charged polynucleotide, suitably DNA, nanostructures into hydrophobic layers, for example phospholipid bilayers, and this issue is expected to increase for nanopores of larger diameter.

There is therefore a need in the field for the provision of membrane-spanning nanopores that fulfil the above criteria of minimum diameter, structural definition and adaptability.

It is also desirable that such pores may be usable not only in biological phospholipid bilayers but also within the membranes which make up synthetic polymer vesicles, membranes or solid state substrates.

SUMMARY OF THE INVENTION

It is an object of the present invention to overcome or alleviate at least one of the above noted drawbacks of prior art systems or to at least provide a useful alternative to related art systems.

In a first aspect the invention relates to a membrane-spanning nanopore, comprising:

-   -   i. at least one scaffold polynucleotide strand;     -   ii. a plurality of staple polynucleotide strands; and     -   iii. at least one hydrophobically-modified polynucleotide         strand, wherein the hydrophobically-modified polynucleotide         strand comprises a polynucleotide strand and a hydrophobic         moiety;     -   wherein each of the plurality of staple polynucleotide strands         hybridises to the or each at least one scaffold polynucleotide         strand to form the three-dimensional structure of the         membrane-spanning nanopore, and wherein the or each at least one         hydrophobically-modified polynucleotide strand hybridises to a         portion of the at least one scaffold polynucleotide strand,     -   the membrane-spanning nanopore defining a central channel with a         minimum internal width of at least about 5 nm.

Typically, the polynucleotide in the or each at least one scaffold strand, each of the plurality of staple polynucleotide strands, and the or each hydrophobically-modified polynucleotide strand comprises DNA. Suitably, the membrane-spanning nanopore of claim 2, wherein the assembly of the nanopore is via DNA origami techniques.

In an embodiment, the minimum internal width of the central channel of the nanopore is from about 5 nm to about 20 nm. Suitably, the minimum internal width of the central channel of the nanopore is from about 5 nm to about 10 nm. In an embodiment, the minimum internal width of the central channel of the nanopore is 7.5 nm.

In an embodiment, the nanopore comprises a membrane-spanning region and at least one cap region. Suitably, the membrane-spanning region is arranged to abut the at least one cap region. In embodiments wherein one cap region is present, the membrane-spanning region is located at one end of the nanopore.

In an embodiment, the membrane-spanning region has a dimension co-axial with the channel of about 1 nm to about 7 nm. Suitably, the membrane-spanning region has a dimension co-axial with the channel of about 3 nm to about 5 nm. In further embodiments, the cap region has a dimension co-axial with the channel of about 20 nm to about 70 nm. Suitably, the cap region has a dimension co-axial with the channel of about 40 nm to about 50 nm.

In an embodiment, the nanopore of the first aspect further comprises one or more adaptor polynucleotide strands, wherein the at least one hydrophobically-modified polynucleotide strand is hybridised to the nanopore via the one or more adaptor polynucleotide strands, the one of more adaptor polynucleotide strands each having a first end and a second end, wherein the first end of the adaptor polynucleotide strand hybridises with the at least one scaffold polynucleotide strand, and the second end of the adaptor polynucleotide strand hybridises with the at least one hydrophobically-modified polynucleotide strand. Suitably, the polynucleotide in the or each adaptor polynucleotide strands is DNA.

In an embodiment, the at least one hydrophobic moiety comprises a lipid. Suitably, the lipid is selected from the group consisting of: sterols; alkylated phenols; flavones; saturated and unsaturated fatty acids; and synthetic lipid molecules (including dodecyl-beta-D-glucoside. Typically, the sterols are selected from the group consisting of: cholesterol; derivatives of cholesterol; phytosterol; ergosterol; and bile acid; the alkylated phenols are selected from the group consisting of: methylated phenols; and tocopherols; the flavones are selected from the group consisting of: flavanone containing compounds; and 6-hydroxyflavone; the saturated and unsaturated fatty acids are selected from the group consisting of: derivatives of lauric acid; oleic acid; linoleic acid; and palmitic acids; and/or the synthetic lipid molecule is dodecyl-beta-D-glucoside.

In an embodiment, the nucleotide sequence of the scaffold strand comprises the DNA sequence of M13mp18 DNA (SEQ ID NO. 1). In further embodiments, the nucleotide sequences of the plurality of staple strands is selected from the group comprising SEQ ID Nos. 2 to 218. In still further embodiments, the nucleotide sequence of the one or more adaptor strands is selected from the group comprising SEQ ID Nos. 219 to 241, and in other embodiments, the sequence of the at least one hydrophobically-modified nucleotide strand is selected from the group comprising SEQ ID No. 242 or 243.

In an embodiment, the nanopore has a cross-section perpendicular to a longitudinal axis of the channel that is quadrilateral in shape. Suitably, the quadrilateral is a square. In another embodiment, the membrane-spanning nanopore is modified, wherein the central channel comprises one or more constrictions.

In a second aspect, the invention provides a membrane comprising one or more of the membrane-spanning nanopores of the first aspect of the invention.

In an embodiment of the second aspect, the membrane comprises a bilayer. The bilayer may be a lipid bilayer. Alternatively, the membrane comprises a semi-fluid membrane formed of polymers. Suitably, the polymer forming the semi-fluid membrane is composed of amphiphilic synthetic block copolymers. Typically, the amphiphilic synthetic block copolymers are composed of hydrophilic copolymer blocks and hydrophobic copolymer blocks. In embodiments, the hydrophilic copolymer blocks are selected from the group consisting of: poly(ethylene glycol) (PEG/PEO); and poly(2-methyloxazoline); and the hydrophobic copolymer blocks are selected from the group consisting of: polydimethylsiloxane (PDMS); poly(caprolactone (PCL); poly(lactide) (PLA); and poly(methyl methacrylate) (PMMA). Suitably, the polymer membrane is composed of the amphiphilic block copolymer poly 2-(methacryloyloxy)ethyl phosphorylcholine-b-disisopropylamino) ethyl methacrylate (PMPC-b-PDPA).

In an embodiment of the second aspect of the invention, the membrane is in the form of a vesicle, a micelle, a planar membrane or a droplet. The membrane may form a droplet interface bilayer or a droplet-droplet bilayer. In a further embodiment, the membrane comprises a solid state substrate.

In a third aspect of the invention, a biological sensor is provided, wherein the biological sensor comprises a membrane of any of the second aspect of the invention and apparatus for measuring an ion flow through one or more membrane-spanning nanopores.

In a fourth aspect, the invention provides a biological sensing device comprising one or more biological sensors of the third aspect of the invention.

In a fifth aspect of the invention, there is provided a method for molecular sensing comprising:

-   -   i. Providing the membrane according to the second aspect of the         invention;     -   ii. Contacting the nanopore with a test substrate and         establishing a flow of ions through the nanopore or an electron         flow across the nanopore; and     -   iii. Measuring the ion flow through the nanopore or electron         flow across one or more of the membrane-spanning nanopores.

In an embodiment of the fifth aspect, the flow of ions is from a first side of the membrane to a second side of the membrane. In a further embodiment, the molecular sensing is analyte detection or characterisation, wherein the change in ion flow or electron flow is indicative of the analyte.

In a further embodiment of the fifth aspect of the invention, the method comprises after step (iii) the further step of determining the presence of the test substrate by a change in ion flow or electron flow through or across the membrane compared to the ion flow or electron flow through or across the membrane when the test substrate is absent.

In an embodiment, the test substrate is a globular protein, a polynucleotide-protein construct, a labelled polynucleotide or a labelled protein.

The sensing apparatus suitable for use in the fifth aspect (and other aspects as appropriate) of the present invention, may comprise a measurement system arranged as disclosed in any of WO 2008/102210, WO 2009/07734, WO 2010/122293, WO 2011/067559 or WO 2014/04443. The sensing apparatus may comprise electrodes arranged on each side of the membrane in order to measure an ion current through an aperture under a potential difference. The electrodes may be connected to an electrical circuit which includes a control circuit arranged to supply a voltage to the electrodes and a measurement circuit arranged to measure the ion flow. A common electrode may be provided to measure ion flow through the apertures between the common electrode and electrodes provided on the opposite side of the membrane.

Fluid chambers provided on either side of the nanopore array may be referred to as the cis and trans chambers. The molecular entity to be determined by the array of nanopores is typically added to the cis chamber comprising the common electrode. Separate trans chambers may be provided on the opposite side of the array, each trans chamber comprising an electrode wherein ion flow through each aperture is measured between an electrode of the trans chamber and the common electrode.

Alternative or additional measurements associated with movement of the molecular entity with respect to the aperture may be carried out, such as measurement of a tunnelling current across the aperture (Ivanov A P et al., Nano Lett. 2011 Jan. 12; 11(1):279-85), or a field effect transistor (FED device, such as disclosed by WO 2005/124888, U.S. Pat. No. 8,828,138, WO 2009/035647, or Xie et al, Nat Nanotechnol. 2011 Dec. 11; 7(2): 119-125. The measurement device may be an FET nanopore device comprising source and drain electrodes to determine the presence or passage of a molecular entity in the apertures. An advantage of employing an FET nanopore device, namely one employing FET measurements across the apertures to measure a local potential or capacitance, or one employing measurement of a tunnelling current across the aperture, is that the measurement signal is very local to a particular aperture and therefore a device comprising a shared trans chamber may be employed. This greatly simplifies the construction of the device without the need to provide separate trans chambers for each aperture, such as one for the measurement of ion flow through the apertures, as described above. As a result, very high densities of apertures in the array may be conveniently provided, for example an array comprising apertures having a pitch of less than 10 nm and a density of 106 apertures/cm².

In a sixth aspect of the invention, a method for molecular gating is provided, the method comprising:

-   -   i. Providing the membrane according to the second aspect of the         invention;     -   ii. Providing at least one biomolecule with a diameter of less         than the minimum internal width of a channel in the nanopore;         and     -   iii. Incubating until the at least one biomolecule has passed         through the nanopore.

In an embodiment of the sixth aspect of the invention, when at least one biomolecule has passed through the nanopore, it is subjected to a physical change that prevents it returning through the nanopore.

In embodiments, the at least one biomolecule is a globular protein, a polynucleotide-protein construct, a labelled polynucleotide or a labelled protein.

A seventh aspect of the invention provides a membrane-spanning nanopore, comprising:

-   -   i. at least one scaffold nucleotide strand comprising at least         one hydrophobic anchor, wherein the hydrophobic anchor comprises         a polynucleotide strand and a hydrophobic moiety; and     -   ii. a plurality of staple polynucleotide strands;     -   wherein each of the plurality of staple polynucleotide strands         hybridises to the or each at least one scaffold polynucleotide         strand to form the three-dimensional structure of the         membrane-spanning nanopore,     -   the membrane-spanning nanopore defining a central channel with a         minimum internal width of at least about 5 nm.

In an embodiment of the seventh aspect, the polynucleotide in the or each at least one scaffold strand, each of the plurality of staple polynucleotide strands, and the or each hydrophobic anchor comprises DNA.

BRIEF DESCRIPTION OF THE DRAWINGS

FIGS. 1A-1C show a design of a large membrane-spanning DNA nanopore designated as NP. (FIG. 1A) The membrane-embedded pore is composed of squarely arranged DNA duplexes which are illustrated as light grey and dark grey cylinders. The latter carry cholesterol lipid anchors (hatched shading) for membrane insertion. Protein trypsin (not shown) can pass via the pore from the cis to the trans side of the membrane. (FIG. 1B) Top-down and side view of the nanopore. (FIG. 1C) Cross-sectional side view to show the geometry of the pore lumen with the dimensions annotated.

FIG. 2 shows a 2D DNA map of the nanopore NP^(−c). The scaffold strand is shown in medium grey and staple strands are in dark grey. The horizontal strands terminating in a dot on the left hand side indicates adaptor strands which hybridize with a portion of their sequence to the cholesterol-modified anchor polynucleotides. 5′ and 3′ termini of DNA strands are represented as a squares and triangles, respectively. The duplexes are numbered at the left.

FIGS. 3A-3C show results of confirming the assembly, purity, dimensions and membrane-interaction properties of DNA nanopores. (FIG. 3A) Gel electrophoretic analysis of scaffold strand (ss) and nanopores NP^(−c), and NP without and with detergent SDS. The position and bp length of the dsDNA markers in kbp is given at the sides of the electropherograms. (FIG. 3B) AFM micrographs (top) and elevation profiles (bottom) of NP^(−c) that assembled on mica into chains. In the micrographs, pores appear as squares due to the compression of the hollow DNA nanostructure by the AFM cantilever. The scale bar for micrographs is 250 and 100 nm for top and bottom images, respectively, and the vertical scale is from 0 to 14 nm for both images. The AFM elevation profiles show the dimensions of the five DNA nanopores in the bottom AFM image. (FIG. 3C) Gel electropherogram of NP and NP^(−c) incubated with no (leftmost lane) or increasing amounts of SUV membrane vesicles ranging from 6.9 to 12.5 nM. The upshifted bands of lipid anchor-bearing NP indicate interaction with bilayer membranes. The interaction does not occur for anchor-free NP^(−c). The right-most band is weaker as half the amount of DNA was loaded. The position of the two dsDNA markers with a length of 10 and 1 kbp are given at the right of the gels.

FIG. 4 shows a size exclusion chromatography trace of an assembly mixture containing the folded DNA nanopore (elution volume of 7.88 mL) and excess staple strands (elution volume of 16.13 mL). Absorption was determined at 260 nm.

FIG. 5 shows and AFM micrograph of cholesterol-free NP^(−c) deposited at a concentration of 2 nM on mica. The DNA nanopores assemble as chains of square-like structures. The square appearance is likely caused by the compression of the hollow nanostructures by the AFM cantilever. The other snake-like objects lacking separate squares are most likely chains of DNA pores but aggregated and altered in conformation by the Ni²⁺ and Mg²⁺ cations used to anchor the DNA to the mica substrate.

FIGS. 6A-6D show rendering images of DNA nanopore NP designed using CaDNAno software and seen from the top (FIG. 6A), bottom (FIG. 6B), side (FIG. 6C), and angular side view (FIG. 6D). The cylinders represent DNA duplexes which are composed of scaffold strand (medium grey) and the staple polynucleotides (dark grey). Light grey strands are adaptor polynucleotides which hybridize with the unpaired single-stranded portion to cholesterol-modified anchor polynucleotides (not shown).

FIGS. 7A-7C shows gel electrophoretic analysis of the interaction between cholesterol-tagged DNA nanopores and lipid vesicles. (FIG. 7A) The DNA nanopores carried the lipid anchor at all 24 possible membrane-spanning duplexes. This corresponds to 12 DNA polynucleotides with the cholesterol TEG modification attached at the 5′ terminus and 12 at the 3′ terminus. (FIGS. 7B, 7C) Gels for pores with (FIG. 7B) 12 lipid anchors at 5′-termini, and (FIG. 7C) 12 lipid anchors at 3′-termini. The lipid concentrations are 0.42 mM (lane 2), 0.58 mM (lane 3), 0.67 mM (lane 4), 0.75 mM (lane 5). The position and bp length of the dsDNA markers are given at the right of the gels.

FIG. 8A-8E show an analysis via single-channel current recordings establishes that NP DNA nanopores span the lipid bilayer. (FIG. 8A) Representative ionic current trace of a single NP pore in 1 M KCl, 10 mM HEPES pH 8.0 and at +20 mV relative to the cis side of the membrane (FIG. 1A). (Fi. 8B) Histogram of channel conductances obtained from 32 independent single-channel recordings at +20 mV to +50 mV. (FIG. 8C) Traces of single DNA pores recorded at voltage ramps running from −100 to +100 mV. The high conductance state is color-coded in dark grey; the lower conductance state is coded in light grey. The traces were selected to show the lower conductance state and hence over-represent its actual frequency of occurrence. (FIG. 8D) IV curve displaying the averages and standard deviation from 10 single-channel current traces. (FIG. 8E) Probabilities for the higher and lower conductance state as a function of the magnitude of positive voltage. Native indicates that the pores were not exposed for at least 10 s to voltages switches while perturbed pores experienced in 1 s intervals at least 8 manually induced voltage switches from +1-40V to +1-100V in 20 mV steps. The probabilities were obtained from the cumulative all-point histogram analysis of multiple traces as described in FIGS. 9A-9B.

FIGS. 9A-9B show all point histograms showing the current distributions of NP pores for the open, high conductance state (solid) and the party-closed lower conductance state (hatched). The current distributions were normalized to the open channel current. The distributions are for (FIG. 9A) native pores held at the recording potential for at least 10 s and (FIG. 9B) perturbed pores that had been exposed to multiple voltage switches at 1 s intervals. The recording potentials are indicated. Histograms for A were obtained by summing up three 15 s segments of independent traces for each voltage while histograms for B were from the sum of each seven 1s segments of independent traces.

FIG. 10 shows Nanopore NP releasing fluorescent probe FAM-PEG³⁵⁰ from giant unilamellar vesicles. Fluorescence images of PEG³⁵⁰-FAM filled vesicles incubated with cholesterol-modified DNA polynucleotides (left) and NP (right).

FIGS. 11A-11C show transport of the protein trypsin through the NP pore. (FIG. 11A) Scheme of the DNA nanopore and its interaction with protein molecules leading to a short encounter without pore translocation (type I) or successful transport (type II). (FIG. 11B) Single-channel current trace recorded in the presence of 1.26 μM trypsin at the cis side leading to blockade of type I and II. Open channel current, I₀, blockade amplitude, A, and dwell time, τ_(off), are defined. (FIG. 11C) Scatter plot showing τ_(off) and A for a single-channel current recording with 1.26 μM trypsin in the cis chamber. Each point in the diagram represents an individual encounter event of protein with the DNA nanopore. The scatter plot comprises 568 data points.

FIG. 12 shows representative tunnelling electron microscope (TEM) images for the DNA nanostructures according to the present invention. Scalebar=50 nm.

FIGS. 13 and 14 show representative tunnelling electron microscope (TEM) images for the DNA nanostructures according to the present invention attached on Single Unicellular Vesicles (SUVs). Scalebar=50 nm.

DETAILED DESCRIPTION

Prior to setting forth the invention, a number of definitions are provided that will assist in the understanding of the invention. All references cited herein are incorporated by reference in their entirety. Unless otherwise defined, all technical and scientific terms used herein have the same meaning as commonly understood by one of ordinary skill in the art to which this invention belongs.

Unless otherwise indicated, the practice of the present invention employs conventional techniques of chemistry, molecular biology, microbiology, recombinant DNA technology, and chemical methods, which are within the capabilities of a person of ordinary skill in the art. Such techniques are also explained in the literature, for example, M. R. Green, J. Sambrook, 2012, Molecular Cloning: A Laboratory Manual, Fourth Edition, Books 1-3, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.; Ausubel, F. M. et al. (1995 and periodic supplements; Current Protocols in Molecular Biology, ch. 9, 13, and 16, John Wiley & Sons, New York, N.Y.); B. Roe, J. Crabtree, and A. Kahn, 1996, DNA Isolation and Sequencing: Essential Techniques, John Wiley & Sons; J. M. Polak and James O′D. McGee, 1990, In Situ Hybridisation: Principles and Practice, Oxford University Press; M. J. Gait (Editor), 1984, Oligonucleotide Synthesis: A Practical Approach, IRL Press; and D. M. J. Lilley and J. E. Dahlberg, 1992, Methods of Enzymology: DNA Structure Part A: Synthesis and Physical Analysis of DNA Methods in Enzymology, Academic Press. Each of these general texts is herein incorporated by reference.

As used herein, the term ‘comprising’ means any of the recited elements are necessarily included and other elements may optionally be included as well. ‘Consisting essentially of’ means any recited elements are necessarily included, elements that would materially affect the basic and novel characteristics of the listed elements are excluded, and other elements may optionally be included. ‘Consisting of’ means that all elements other than those listed are excluded. Embodiments defined by each of these terms are within the scope of this invention.

The term ‘membrane’ as used herein is an enclosing or separating selectively-permeable boundary, partition, barrier or film. The membrane has two sides or surfaces which may be named the cis and trans side respectively. The membrane is thin (i.e. has a thickness substantially less than its width and length) allowing it to be spanned by the nanopore. In the context of the present invention, the membrane thickness is the typically in the nanometre (10⁻⁹ metre) range. The arrangement of the membrane is not limited and may in any form, for example, a liposome, vesicle or as a planar or a non-planar sheet. Specific examples of membranes useful in the present invention include lipid bilayers, polymeric films, or solid state substrates.

The term ‘solid state membrane’ or ‘solid state substrate’ as used herein refers to a membrane formed from a solid state substance in which one or more apertures are provided. One or more nanopores may be positioned within the respective one or more apertures disclosed for example in U.S. Pat. No. 8,828,211, hereby incorporated by reference. The solid state membrane may comprise either or both of organic and inorganic materials, including, but not limited to, microelectronic materials, whether electrically conducting, electrically semiconducting, or electrically insulating, including materials such as II-IV and III-V materials, oxides and nitrides, such as silicon nitride, Al₂O₃, and SiO₂, Si, MoS₂, solid state organic and inorganic polymers such as polyamide, plastics such as Teflon®, or elastomers such as two-component addition-cure silicone rubber, and glasses. A membrane may be formed from monatomic layers, such as graphene, or layers that are only a few atoms thick such as those disclosed in U.S. Pat. No. 8,698,481, and U.S. Patent Application Publication 2014/174927, both hereby incorporated by reference. More than one layer of material can be included, such as more than one graphene layer, as disclosed in US Patent Application Publication 2013/309776, incorporated herein by reference. Suitable silicon nitride membranes are disclosed in U.S. Pat. No. 6,627,067, and the membrane may be chemically functionalized, such as disclosed in U.S. Patent Application Publication 2011/053284, both hereby incorporated by reference. Such a structure is disclosed for example in U.S. Pat. No. 8,828,211, hereby incorporated by reference. The internal walls of the apertures may be coated with a functionalised coating, such as disclosed in published application WO 2009/020682. The one or more apertures may be hydrophobic or provided with a hydrophobic coating to assist the provision of the one or more nanopores in the respective one or more apertures. Suitable methods for providing apertures in solid state substrates are disclosed in published applications WO 03003446 and WO 2016/187519.

The term ‘nucleic acid’ as used herein, is a single or double stranded covalently-linked sequence of nucleotides in which the 3′ and 5′ ends on each nucleotide are joined by phosphodiester bonds. The polynucleotide may be made up of deoxyribonucleotide bases or ribonucleotide bases. Nucleic acids may include DNA and RNA, and are typically manufactured synthetically, but may also be isolated from natural sources. Nucleic acids may further include modified DNA or RNA, for example DNA or RNA that has been methylated or that has been subject to chemical modification, for example 5′-capping with 7-methylguanosine, 3′-processing such as cleavage and polyadenylation, and splicing, or labelling with fluorophores or other compounds. Nucleic acids may also include synthetic nucleic acids (XNA), such as hexitol nucleic acid (HNA), cyclohexene nucleic acid (CeNA), threose nucleic acid (TNA), glycerol nucleic acid (GNA), locked nucleic acid (LNA) and peptide nucleic acid (PNA). Hence, where the terms DNA′ and ‘RNA’ are used herein it should be understood that these terms are not limited to only include naturally occurring nucleotides. Sizes of nucleic acids, also referred to herein as ‘polynucleotides’ are typically expressed as the number of base pairs (bp) for double stranded polynucleotides, or in the case of single stranded polynucleotides as the number of nucleotides (nt). One thousand bp or nt equal a kilobase (kb). Polynucleotides of less than around 100 nucleotides in length are typically called ‘oligonucleotides’.

As used herein, the terms ‘3′’ (‘3 prime’) and ‘5′’ (‘5 prime’) take their usual meanings in the art, i.e. to distinguish the ends of polynucleotides. A polynucleotide has a 5′ and a 3′ end and polynucleotide sequences are conventionally written in a 5′ to 3′ direction. The term ‘complements of a polynucleotide molecule’ denotes a polynucleotide molecule having a complementary base sequence and reverse orientation as compared to a reference sequence.

The term ‘duplex’ is used herein refers to double-stranded DNA, meaning that the nucleotides of two complimentary DNA sequences have bonded together and then coiled to form a double helix.

According to the present invention, homology to the nucleic acid sequences described herein is not limited simply to 100% sequence identity. Many nucleic acid sequences can demonstrate biochemical equivalence to each other despite having apparently low sequence identity. In the present invention homologous nucleic acid sequences are considered to be those that will hybridise to each other under conditions of low stringency (Sambrook J. et al, supra).

As used herein, the term ‘nanostructure’ refers to a predesigned two or three dimensional molecular structure typically comprised from a biopolymer, suitably a naturally or non-naturally occurring nucleic acid, which structure has at least one dimension or an aspect of its geometry that is within the nanoscale (i.e. 10⁻⁹ metres). Nanoscale structures suitably have dimensions or geometry of less than around 100 nm, typically less than 50 nm, and most suitably less than 20 nm. Nanoscale structures suitably possess dimensions or geometry greater than around 0.1 nm, typically greater than around 1 nm, and optionally greater than around 2 nm. Assembly of nucleic acid nanostructures may occur spontaneously in solution, or may require presence of additional co-factors including, but not limited to, nucleic acid scaffolds, nucleic acid aptamers, nucleic acid staples, co-enzymes, and molecular chaperones. Where desired nanostructures result from one or more predesigned spontaneously self-folding nucleic acid molecules, such as DNA, this is typically referred to as nucleic acid ‘origami’. Exemplary three dimensional DNA nanostructures may comprise nanobarrels; nanorafts, which are typically rectangular, polygonal, circular, or ellipsoid substantially planar nanostructures; nanospheres and regular or irregular polyhedral nanostructures, including stellated polyhedral nanostructures. Exemplary two dimensional DNA nanostructures may comprise nanodiscs or nanoplates. Rational design and folding of DNA to create two dimensional or three dimensional nanoscale structures and shapes is known in the art (e.g. Rothemund (2006) Nature 440, 297-302).

The nucleic acid sequences that form the nanostructures will typically be manufactured synthetically, although they may also be obtained by conventional recombinant nucleic acid techniques. DNA constructs comprising the required sequences may be comprised within vectors grown within a microbial host organism (such as E. coli). This would allow for large quantities of the DNA to be prepared within a bioreactor and then harvested using conventional techniques. The vectors may be isolated, purified to remove extraneous material, with the desired DNA sequences excised by restriction endonucleases and isolated, such as by using chromatographic or electrophoretic separation.

The term ‘amino acid’ in the context of the present invention is used in its broadest sense and is meant to include naturally occurring L α-amino acids or residues. The commonly used one and three letter abbreviations for naturally occurring amino acids are used herein: A=Ala; C=Cys; D=Asp; E=Glu; F=Phe; G=Gly; H=His; I=Ile; K=Lys; L=Leu; M=Met; N=Asn; P=Pro; Q=Gln; R=Arg; S=Ser; T=Thr; V=Val; W=Trp; and Y=Tyr (Lehninger, A. L., (1975) Biochemistry, 2d ed., pp. 71-92, Worth Publishers, New York). The general term ‘amino acid’ further includes D-amino acids, retro-inverso amino acids as well as chemically modified amino acids such as amino acid analogues, naturally occurring amino acids that are not usually incorporated into proteins such as norleucine, and chemically synthesised compounds having properties known in the art to be characteristic of an amino acid, such as β-amino acids. For example, analogues or mimetics of phenylalanine or proline, which allow the same conformational restriction of the peptide compounds as do natural Phe or Pro, are included within the definition of amino acid. Such analogues and mimetics are referred to herein as ‘functional equivalents’ of the respective amino acid. Other examples of amino acids are listed by Roberts and Vellaccio, The Peptides: Analysis, Synthesis, Biology, Gross and Meiehofer, eds., Vol. 5 p. 341, Academic Press, Inc., N.Y. 1983, which is incorporated herein by reference.

A ‘polypeptide’ is a polymer of amino acid residues joined by peptide bonds, whether produced naturally or in vitro by synthetic means. Polypeptides of less than around 12 amino acid residues in length are typically referred to as ‘peptides’ and those between about 12 and about 30 amino acid residues in length may be referred to as ‘oligopeptides’. The term ‘polypeptide’ as used herein denotes the product of a naturally occurring polypeptide, precursor form or proprotein. Polypeptides can also undergo maturation or post-translational modification processes that may include, but are not limited to: glycosylation, proteolytic cleavage, lipidization, signal peptide cleavage, propeptide cleavage, phosphorylation, and such like. The term ‘protein’ is used herein to refer to a macromolecule comprising one or more polypeptide chains.

The term ‘folded protein’ as used herein refers to a protein that has acquired some three-dimensional shape after translation of the polypeptide chain from which it is formed (the primary structure). The term may refer to the secondary structure of the protein which is typically the first stage of the folding process where local three-dimensional structures are formed, for example, alpha helices or beta sheets. The term may more typically refer to the tertiary structure of a protein where the secondary structures of the protein have folded to stabilise the structure through hydrophobic or covalent interactions. The term also encompasses proteins having a quaternary structure where one or more protein subunits are assembled. As appropriate, the folded protein may also be termed the ‘native’ protein structure, and may be the form of the protein that exhibits its biological function.

The term ‘interior width’ when used herein refers to the straight distance spanning the interior of the channel from an interior face of one wall to an interior face of an opposing wall in a plane perpendicular to the longitudinal axis of the channel. The interior width of the channel may be constant along its longitudinal axis or it may vary. The ‘minimum interior width’ is the minimum interior width along the longitudinal axis of the channel between an entrance and an exit of the channel. The minimum interior width of a channel defines the maximum size of an object that may pass through the channel.

As used herein the term ‘hydrophobic’ refers to a molecule having apolar character including organic molecules and polymers. Examples are saturated or unsaturated hydrocarbons. The molecule may have amphipathic properties.

As used herein, the term ‘hydrophobically-modified’ relates to the modification (joining, bonding or otherwise linking) of a polynucleotide strand with one or more hydrophobic moieties. A ‘hydrophobic moiety’ as defined herein is a hydrophobic organic molecule. The hydrophobic moiety may be any moiety comprising non-polar or low polarity aliphatic, aliphatic-aromatic or aromatic chains. Suitably, the hydrophobic moieties utilised in the present invention encompass molecules such as long chain carbocyclic molecules, polymers, block co-polymers, and lipids. The term ‘lipids’ as defined herein relates to fatty acids and their derivatives (including tri-, di-, monoglycerides, and phospholipids), as well as sterol-containing metabolites such as cholesterol. The hydrophobic moieties comprised within the embodiments of the present invention are capable of forming non-covalent attractive interactions with phospholipid bilayers, such as the lipid-based membranes of cells and act as membrane anchors for the nanopore. According to certain embodiments of the present invention suitable hydrophobic moieties, such as lipid molecules, possessing membrane anchoring properties may include sterols (including cholesterol, derivatives of cholesterol, phytosterol, ergosterol and bile acid), alkylated phenols (including methylated phenols and tocopherols), flavones (including flavanone containing compounds such as 6-hydroxyflavone), saturated and unsaturated fatty acids (including derivatives such as lauric, oleic, linoleic and palmitic acids), and synthetic lipid molecules (including dodecyl-beta-D-glucoside). The anchors for the polymer membrane may be the same as for lipid bilayers or they may be different. The specific hydrophobic moiety anchor may be selected based on the binding performance of the membrane chosen.

The inventors have identified a structurally-defined wide-channel organic nanopore. The structure of the nanopore renders it suitable for a number of uses. One such use is biomolecule sensing applications, in particular protein sensing applications where the folded protein may pass, bind or lodge within the pore. Modified versions of the nanopore may serve to further enhance the suitability of the channel for a particular folded protein or other biomolecule. Biomolecule sensing may be enhanced by the installation of a molecular receptor within the pore lumen or channel.

The wide channel DNA nanopore according to the present invention offers an advantage over existing protein pores such as ClyA or alpha-hemolysin, and known DNA nanopores. The nanostructure provides a wide, stable membrane-spanning channel that allows folded proteins, typically proteins that have a three-dimensional tertiary or quaternary structure, to pass, bind or lodge within. Passing or binding a folded protein in the channel of a nanopore leads to an improved read-out signal through more effective blockade of the ion flow through the open channel, as compared to binding of the protein at the mouth of the channel.

While DNA pores are, by their nature, more negatively charged than usual protein pores. Potential electrostatic repulsion of cargo, if it were to occur, could be compensated by higher ionic strength or charge-neutralized DNA (Burns J., Stulz E., Howorka S. Self-assembled DNA nanopores that span lipid bilayers. Nano Lett. 13, 2351-2356 (2013)).

The wide opening of the channel of the nanopore also facilitates the translocation of folded proteins and large biomolecules through a polymer membrane leading to potential applications of the nanopores in molecular gating of biomolecules. Recently developed synthetic membrane-spanning DNA nanopores provide a new and potentially generic route for controlled transport across membranes (see, for example, Douglas S. M., Marblestone A. H., Teerapittayanon S., Vazquez A., Church G. M., Shih W. M. Nucleic Acids Res. 37, 5001-5006 (2009); Zheng J., et al. Nature 461, 74-77 (2009); Fu J., et al. Nat. Nanotechnol. 9, 531-536 (2014); Burns J. R., et al. Angew. Chem. Int. Ed. 52, 12069-12072 (2013); and Seifert A., Göpfrich K., Burns J. R., Fertig N., Keyser U. F., Howorka S. ACS Nano 9, 1117-1126 (2015)).

The ability to create nanopores with controlled pore sizes and insert them into natural or synthetic membranes allows the construction of customised selectively-permeable membranes where control over the lumen dimensions, and optionally, other features of the pores enables control over which biomolecules are able to pass through these membranes. Of particular utility in the present invention is the increased size of the nanopores of the present invention that allows large biomolecules, including large globular proteins, to pass through such membranes which has not been possible to date. This has potential utility in medicine alongside other uses in the field of biology. For example, there are several large i.e. 10-30 nm diameter membrane pores that are either produced by immune cells to kill bacteria, such as in the ‘complement system’ (a part of the innate immune system that enhances (complements) the ability of antibodies and phagocytic cells to clear microbes and damaged cells from an organism, promotes inflammation, and attacks the pathogen's plasma membrane). In particular to the C9 protein forms a large pore that together with other accessory proteins forms the membrane attack complex. Membrane portion is also used by pathogenic bacteria to attack and kill eukaryotic host cells. Examples of these bacterial pores are Cholesterol-dependent Cytolysin which includes perfrinoglysin or listeriolysin O.

As an example, vesicles formed of natural or synthetic polymers comprising nanopores according to the present invention may be used as nanoreactors. It is envisaged that substrate proteins or biomolecules may enter the vesicle interior through the nanopore where encapsulated enzymes in the interior of the vesicle enable a desired reaction to take place within the vesicle. As an example, encapsulated protease enzymes, such as trypsin may be retained inside a vesicle where the relatively large and controllable size of the nanopores could allow some substrate proteins but not others to enter for degradation, thereby protecting particular proteins from indiscriminate digestion. Similarly, control over the release of cargo contained within similar vesicles, such as protein or peptide based pharmaceutical agents would be possible with membranes with nanopores which allow passage of the cargo.

According to an embodiment of the present invention, there is provided novel nucleic acid nanopores that have a central channel or lumen with a relatively large minimum diameter (for example, greater than about 5 nm) surrounded and defined by a generally elongate cylindrical pore wall. As described herein, the nanopores are structurally defined and are tunable to permit adaption to different pore channel sizes. The pores may be composed of any suitable nucleic acids. Nanopores formed from one or more DNA duplexes are particularly suitable as they are an excellent construction material for rationally designing nanoscale architectures of defined size (Langecker M., et al. Science 338, 932-936 (2012); Burns J., Stulz E., Howorka S. Nano Lett. 13, 2351-2356 (2013); Burns J. R., Al-Juffali N., Janes S. M., Howorka S. Angew. Chem. Int. Ed. 53, 12466-12470 (2014); Burns J. R., Seifert A., Fertig N., Howorka S. Nat. Nanotechnol. 11, 152-156 (2016)).

The nanopore may be designed to have any suitable shape, although generally it has an elongated cylindrical shape. The dimensions of the nanopore are determined to a degree by the length of the scaffold strand used, the number of layers of nucleic acid duplex used to line the pore and the height of the pore. For example, for a three-layer square nanopore design based on the use of the scaffold strand M13mp18 that is 7249 bases long, a pore may be designed with a minimum internal width or opening of approximately 40 nm, and a diagonal width of 57 nm. In an alternative possible design, using the same scaffold strand M13mp18, a nanopore having a maximum length of 310 nm although the minimum internal width in only 2.5 nm, and therefore outside the scope of this invention.

The nanopore according to an embodiment of the present invention comprises two regions: a cap region and a membrane-spanning region. The membrane-spanning region is defined as the portion of the nanopore located within the plane of the membrane, and the cap region being the portion of the nanopore attached to the membrane-spanning region and extending away from the surface of the membrane. The nanopore may have a cap region on one side of the membrane only, or alternatively, have two cap regions, one on each side of the membrane. When there is more than one cap region, these may be the same as each other or they may be different. Suitably, the nanopore has one cap region on one side of the membrane forming an entrance to the nanopore.

The cap region may have dimensions of any suitable size. While it is possible for the cap region to extend only negligibly from the membrane surface, typically, the cap region has a height extending from the membrane of at least 5 nm as measured by the perpendicular distance from the membrane surface to the top of the pore wall. Suitably, the cap region may have a height of at least 10 nm, 15 nm, 20 nm, 25 nm, 30 nm, 35 nm, 40 nm, 45 nm or 50 nm or above. Suitably, the height of the cap region is at most 100 nm, 90 nm, 80 nm, 70 nm, 60 nm, or 50 nm or below. The height of the cap region may be determined by the length of the scaffold polynucleotide used, and the number of layers of polynucleotide duplexes that form the pore. For example, according to calculations using computer software (CaDNAno software, available at http://www.cadnano.org), for a square cross-section DNA pore using M13mp18 scaffold strand and with a minimum interior width of the pore of 20 nm, the maximum height of the pore is around 37 nm when the pore wall is two duplexes thick; 20 nm when the pore wall is three duplexes thick; and 13 nm when the pore is four duplexes thick.

The membrane-spanning region may have dimensions of any suitable size. Typically, the membrane-spanning region has a height that approximately matches the thickness of the membrane in which it resides. The thickness of biological lipid bilayer membranes can range from around 3.5 to 10 nm. The thickness of membrane composed of amphiphilic synthetic block copolymers shows a wider range from 5 to 50 nm (C. LoPresti, H. Lomas, M. Massignani, T. Smart, G. Battaglia, J. Mater. Chem. 2009, 19, 3576-3590). Therefore suitably, the membrane-spanning region may have a height of at least around 3.5 nm, although it may be possible to have a membrane-spanning region with a height as low as 3 nm, 2.5 nm, 2 nm 1.5 nm or 1.0 nm or less. Suitably, the membrane-spanning region may have a height of at least 5 nm. The membrane-spanning region may have a height of at most 100 nm, 90 nm, 80 nm, 70 nm, 60 nm, 50 nm, 40 nm, 30 nm, 20 nm or 10 nm or less. Suitably the membrane-spanning region has a maximum height of 50 nm for synthetic polymer layers, and a maximum height of 10 nm for lipid bilayers.

The channel or lumen that passes through the nanopore has a cross-sectional profile parallel to the membrane and perpendicular to a longitudinal axis of the channel. This cross-sectional profile may be of any shape and dimensions. The shape and dimensions of the channel may be consistent for its entire length or may vary. Suitably, the channel has a consistent cross-sectional profile and size for its entire length. Suitably, the cross-sectional profile of the channel is a quadrilateral, typically a square, or at least generally circular. The minimum opening or width of the channel in this cross-section is suitable to allow access for a folded protein. Typically, the minimum opening or width of the channel is at least 5 nm, 6 nm, 6.5 nm, 7 nm, 7.5 nm, 8 nm, 8.5 nm, 9 nm, 9.5 nm, 10 nm, 11 nm, 12 nm, 13 nm, 14 nm, 15 nm, 16 nm, 17 nm, 18 nm, 19 nm, 20 nm, 25 nm or more. Suitably the opening is between around 5 nm and around 20 nm. Typically the opening is around 5 nm and around 10 nm. The maximum opening of the channel is limited only by the need to maintain structural integrity of the pore and to obtain an electrical read-out when a molecule of interest passes through. Suitably, the maximum width or opening of the channel is 50 nm, 45 nm, 40 nm, 35 nm, 30 nm, 25 nm, 20 nm, 18 nm, 15 nm, 12 nm, or 10 nm. Suitably, the cross-sectional area of the minimum opening of the channel is at least 20 nm², 25 nm², 30 nm², 35 nm², 40 nm², 45 nm², 50 nm², 60 nm², 70 nm², 80 nm², 90 nm² or 100 nm² or more. Suitably, the cross-sectional area of the minimum opening of the channel is at most 200 nm², 180 nm², 160 nm², 140 nm², 120 nm², 100 nm², 90 nm², 80 nm², 70 nm², 60 nm², 50 nm², 40 nm², 30 nm², 20 nm² or 15 nm² or less.

To achieve optimum performance, the variation in the channel size should be minimized from pore to pore. Even small variation in surface area can lead to a significant discrepancy in ion flow through a given pore, both in the open state (devoid of any target analyte), or in the bound state (with target analyte present in, or proximate to, the channel). Such variation in ion flow leads to a lower signal to noise ratio in the electrical read-out thereby reducing the sensitivity of detection. The nanopores according to the present invention have low variability in pore size (see FIGS. 12 to 14 ).

The nanopore may be modified in order to provide one or more constrictions. The constriction may restrict the channel width. The constriction may restrict the channel width from between about 0.5 nm up to the width of the channel. A constriction is a narrowing of the channel width and more than one constriction may be provided along the length of the channel. The channel may be provided for example with two such constrictions. The constrictions may be spaced apart. The constrictions may be spaced apart from each other by 1 nm or more, for example spaced apart by a value between 1 nm and 20 nm, more particularly 5 and 10 nm. The modification may be chemical. A modification may be made after assembly of the nanopore. Alternatively modification may be made to the scaffold and/or staple strands prior to assembly of the nanopore. Examples of chemical modifications to protein nanopores in order to narrow the channel diameter are described in PCT/GB2017/050961.

Suitably, the nanopores of the present invention are assembled via the ‘scaffold-and-staple’ approach. In this important route to nucleic acid nanostructure, in particular, DNA nanostructures, DNA is utilized as a building material in order to make nanoscale three dimensional shapes. Assembly of these complex nanostructures from a plurality of un-hybridized linear molecules is typically referred to as ‘DNA origami’, although the technique is equally applicable to other nucleic acids.

The DNA origami process generally involves the folding of the one or more elongate, ‘scaffold’ DNA strands into a particular shape using a plurality of rationally designed ‘staple’ DNA strands. The scaffold strand can have any sufficiently non-repetitive sequence. There are many DNA sequences that are suitable for use as a scaffold sequences. Commercially available examples include single stranded scaffold DNA sequences named according to their base length, for example, P7249 (base sequence as in M13mp18; 7249 bases); type p7560 (7560 bases); p8064 (8064 bases) available from, for example, Eurofins Genomics (https://www.eurofinsgenomics.eu/en/dna-rna-oligonucleotides/oligo-design-more/dna-origami/scaffold-dna.aspx; accessed 13 Jul. 2017). The sequences of the staple strands are designed such that they hybridize to particular defined portions of the scaffold strands and, in doing so, force the scaffold strands to assume a particular configuration. Methods useful in the making of DNA origami structures can be found, for example, in Rothemund, P. W., Nature 440:297-302 (2006); Douglas et al, Nature 459:414-418 (2009); Dietz et al, Science 325:725-730 (2009); and U.S. Pat. App. Pub. Nos. 2007/0117109, 2008/0287668, 2010/0069621 and 2010/0216978, each of which is incorporated by reference in its entirety. Staple sequence design can be facilitated using, for example, CaDNAno software, available at http://www.cadnano.org.

In the context of the present invention, the sequences of the staple strands are selected such that the polynucleotide, suitably DNA, nanostructure assumes a shape or configuration that corresponds to a membrane-spanning nanostructure. In some embodiments, the staple strands of the DNA nanostructure are selected such that the DNA nanostructure is substantially channel- or tube-shaped. In such embodiments, the inner surface of the DNA nanostructure is the surface on the inside of the tube—i.e. within the interior of the membrane spanning channel—while the outer surface of the DNA nanostructure is the outside of the tube.

In some embodiments, the scaffold strands of the DNA nanostructure are selected such that the membrane-spanning nanostructure has a first DNA duplex domain and second or further DNA duplex domains, wherein a first end of a first domain is attached to a first end of a second or further domain by one or more single-stranded DNA hinges or cross-over sequences, or simply ‘cross-overs’. (Burns J. R., et al. Angew. Chem. Int. Ed. 52, 12069-12072 (2013); Burns J. R., Al-Juffali N., Janes S. M., Howorka S. Angew. Chem. Int. Ed. 53, 12466-12470 (2014); and Yang Y., Zhao Z., Zhang F., Nangreave J., Liu Y., Yan H. Nano Lett. 13, 1862-1866 (2013)). The cross-overs can be simple single-stranded DNA loops that connect two DNA duplexes such as at the termini of the two duplexes. Cross-over also includes structures where two DNA duplexes are interlinked by two strands at the same site. This type of cross-over is called a Holliday junction. A Holliday junction is a branched nucleic acid structure that contains four double-stranded arms joined together. A Holliday junction may be in the form of a cross, however in the case of DNA nanostructures, the four arms of the cross are more parallel whereby each two arms on one side form a duplex. A reference for cross-overs in DNA nanotech structures is Seeman, N. C., Annu. Rev. Biochem. 79, 65-87 (2010).

The nanopore of the present invention may comprise one or more hydrophobic moieties that act as anchors to attach or connect or anchor the hydrophilic DNA nanopore to the generally hydrophobic membrane (lipid bilayer, polymer or solid state). The hydrophobic anchors are attached to the pore. Suitably attachment is via polynucleotides, suitably DNA polynucleotide strands that carry the hydrophobic moiety, suitably a lipid such as cholesterol, at the 5′ or 3′ terminus. These hydrophobically-modified anchor strands hybridize via ‘adaptor’ polynucleotide strands to corresponding sections of the polynucleotide sequence forming the scaffold section of the pore. Alternatively, the hydrophobic anchors are assembled with the pore using hydrophobically-modified polynucleotides. The number of hydrophobically-modified anchors on a single pore is not limited and may number 20, 19, 18, 17, 16, 15, 14, 13, 12, 11, 10, 9, 8, 7, 6, 5, 4, 3, 2 or 1. Cholesterol has been found to be a particularly suitable hydrophobic moiety for use as an anchor in the present invention. The use of other lipids as anchors is contemplated, although it may be expected that there is a particular preference for a particular hydrophobic moiety, and a given number of hydrophobic anchors, for a given membrane.

The membrane in which the nanopore of the present invention may be inserted may be of any suitable type. Depending on the intended use, the membrane may a lipid bilayer or a polymer sheet or film or a solid state substrate. In solid state membranes, the substrate may already comprise apertures in which the nanopore sits thereby adapting the interior dimensions and channel width of the aperture. The membrane is typically hydrophobic to promote anchoring by the hydrophobic anchors.

Lipid bilayers are ubiquitous in biological organisms. It is envisaged that nanopores according to the present invention may be inserted into a lipid membrane of a target cell or vesicle to facilitate translocation of specific folded proteins across the membrane. The specificity of the translocation to certain folded proteins may be controlled through variation of the size of the channel in the nanopore.

Proprietary and non-proprietary synthetic polymer films or sheets are widely used in ‘chip-based’ nanopore sequencing and analytical applications such as the MinION® system sold by Oxford Nanopore Technologies®; the GS FLX+® and the GS Junior® System sold by Roche®; the HiSeg®, Genome Analyzer IIx®, MiSeq® and the HiScanSQ® systems sold by Illumina®; the Ion PGM® System and the Ion Proton System® sold by Life Technologies; the CEQ® system sold by Beckman Coulter®; and the PacBio RS® and the SMRT® system sold by Pacific Biosciences®. The ability of nanopores to insert into polymer membranes of this type would allow these systems to be adapted for folded protein sensing applications.

The polymer membrane may be formed of any suitable material. Typically, synthetic membranes are composed of amphiphilic synthetic block copolymers. Examples of hydrophilic block copolymers are poly(ethylene glycol) (PEG/PEO) or poly(2-methyloxazoline), while examples of hydrophobic blocks are polydimethylsiloxane (PDMS), poly(caprolactone) (PCL), poly(lactide) (PLA), or poly(methyl methacrylate) (PMMA). In embodiments, the polymer membrane used may be formed from the amphiphilic block copolymer poly 2-(methacryloyloxy)ethyl phosphorylcholine-b-disisopropylamino) ethyl methacrylate (PMPC-b-PDPA). DNA nanopores may be inserted into the walls of such polymersomes through incubation. Without wishing to be bound by theory, it is believed that the process of insertion likely involves a first step of membrane tethering, followed in a second step by vertical re-orientation of the DNA pore to achieve complete insertion. This however requires cholesterol anchors to be comprised within the pores, without which insertion does not take place. One exemplary embodiment is described in Example 2 below.

The ability to adapt the nanopores of the present invention such that they may be inserted into membranes of varying thickness is a significant advantage.

The molecular sensors and devices disclosed herein may be used for the detection or characterization of an analyte. Whilst the channel dimensions of the nanopores disclosed herein are particularly suitable for the detection of larger analytes such as unfolded or globular proteins, the nanopores are also suitable for the detection of other analytes. The nanopores may be modified to reduce the width of the channel making the nanopore suitable for the detection of analytes having a smaller width. However nanopores without the modified channel may also be used to detect analytes having a smaller width. Conversely modified nanopores may be used to determine analytes having a larger width than the width of the channel or constriction.

The analyte may be caused to fully or partially translocate the channel. The analyte may for example be held or lodged within the channel or channel entrance, for example at a constriction. Measurement of a signal, for example the change in ion flow during translocation may be used to detect or characterize the analyte. Alternatively the analyte may be caused to pass across the entrance of the nanopore channel in order to detect or characterize it.

Examples of analytes that may be detected or characterized are folded or unfolded proteins, DNA-protein constructs such as nucleosomes and polynucleotides such as deoxyribonucleic acid (DNA) or ribonucleic acid (RNA), polysaccharides and synthetic polymers. The target polynucleotide can comprise one strand of RNA hybridized to one strand of DNA. The polynucleotide may be any synthetic nucleic acid known in the art, such as peptide nucleic acid (PNA), glycerol nucleic acid (GNA), threose nucleic acid (TNA), locked nucleic acid (LNA) or other synthetic polymers with nucleotide side chains. The polynucleotide can be single or double stranded or have a tertiary or quaternary structure. The proteins or nucleic acids can be labelled with a detectable label. The label may be optically detectable such as a fluorophore. The analyte may be an enzyme and the sensor may be used to determine a conformational change in the enzyme.

The presence or absence of the analyte may be detected. Alternatively the analyte may be characterized, for example in the case of a nucleic acid, the sequence of the nucleotides may be determined from characteristic disruptions in the measured signal over time. In the case of proteins, aspects of the protein structure may be determined. The protein may be unfolded prior to detection using the nanopore. An example of such is disclosed in PCT/US2013/026414.

The invention is further illustrated by the following non-limiting examples.

EXAMPLES Materials and Methods

Native and cholesterol-labeled DNA polynucleotides with a tri(ethylene glycol)(TEG) linker were purchased from Integrated DNA Technologies (Leuven, Belgium) or ATDbio (Southampton, United Kingdom) on a 1 μmole scale with desalting or HPLC purification, respectively. 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE), 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) and 1,2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC) was procured from Avanti Polar Lipids (Alabaster, Ala.). M13mp18 DNA was from New England Biolabs (Ipswich, United Kingdom). PEG³⁵⁰-FAM was procured form Chem Quest (United Kingdom). All other reagents and solvents were purchased from Sigma-Aldrich unless stated.

Nanopore Design

The DNA origami nanopores were designed using the square-lattice version of the CaDNAno software (Douglas S. M., Marblestone A. H., Teerapittayanon S., Vazquez A., Church G. M., Shih W. M. Nucleic Acids Res. 37, 5001-5006 (2009)). To assess rigidity in the structural design, several cycles of strand routing with CaDNAno and CanDo (Castro C. E., et al. Nat. Methods 8, 221-229 (2011)) modeling were conducted. The 7249 nt-long single stranded M13mp18 DNA was selected as the scaffold strand. The rendering of DNA nanopore and the 2D DNA map highlighting the scaffold in medium grey and staple strands in dark grey as shown in FIGS. 6 and 2 , respectively. In the design, lipid anchors are attached to the pore via DNA polynucleotides that carry cholesterol at the 5′ or 3′ terminus. These cholesterol-modified anchor strands hybridize via adaptor polynucleotides to the pore; the adapter-mediated binding enables limitation of the number of expensive cholesterol-modified polynucleotides to two. The DNA sequences of staple strands, adaptor strands, and cholesterol-modified anchor strands are provided in Table 1.

Assembly

DNA nanopores were formed by first annealing NP^(−c) in a one-pot reaction containing 1×TAE buffer, supplemented with 14 mM MgCl₂, and a mixture of M13mp18 scaffold and staples at final concentrations of 4.2 nM and 100 nM, respectively. Assembly was conducted using a 7 day-long protocol involving a first annealing phase from 80° C. to 60° C. at a cooling rate of 1° C. per 5 min, and a second phase from 60° C. to 20° C. at a rate of 1° C. per 300 min. To form nanopore NP with cholesterol lipid anchors, NP^(−c) that underwent purification with size-exclusion chromatography (see below) was mixed with cholesterol-modified anchor polynucleotides (1.1 eq. strand per binding site at the pore, up to 24 sites) and incubated at 30° C. for 12 h.

Agarose Gel Electrophoresis of DNA Nanopores without and with Vesicles

The assembly products were analyzed using 1.5% agarose gel electrophoresis in standard 1×TAE buffer, optionally supplemented with 0.015% SDS. DNA pore samples (10 μL) were mixed with 6× gel loading buffer (2 μL) and then loaded into the wells. Gels were run at 70 V for 1 h at 8° C. A 1000-base-pair marker (New England Biolabs) was used as the reference standard. DNA bands were visualized by staining with ethidium bromide solution and ultraviolet illumination. SDS containing gels were washed with deionized water for 20 min prior to staining.

To analyze the interaction of NPs with membranes, small unilamellar vesicles (SUVs) were formed. Chloroform solutions of DOPE (0.3 mmol, 22.3 μL) and DOPC (0.7 mmol, 110 μL) were mixed, and added to an oven-dried round bottom flask (10 mL), followed by removal of the solvent under vacuum using a rotary evaporator for 20 min. To form vesicles, a solution of 0.3 M KCl, 15 mM Tris, pH 8.0 (1 mL) was added, and the suspension was sonicated for 20 min at RT. SUV preparations were stored at 4° C. and used within one week. Before experimentation, the SUV solution was vortexed for 2 s. For agarose gel electrophoretic analysis of nanopores with SUVs, the same gel conditions described above were used, except that SDS was omitted and gels were run at 40 V. Pores (15 μL, 1 μM, 0.3 M KCl, 15 mM Tris, pH 8.0) were incubated with SUVs (15 μL, 1 mM, 0.3 M KCl, 15 mM Tris, pH 8.0) for 30 min at 37° C. To the mixture, blue loading dye (6×, no SDS, 10 μL) was added and loaded onto the gel (30 μL).

Purification

The assembled nanopores were purified from excess staples via size exclusion chromatography (SEC) using an AKTA purifier 100/10 fitted with a Superdex 200 10/300 GL column (GE Healthcare), using a flow rate of 0.5 mL per min at 8° C. Elution was monitored via UV-vis absorption at 260, 280 and 295 nm, and fractions containing the folded DNA pore were pooled.

Atomic Force Microscopy

DNA origami pore NP^(−c) was imaged using tapping mode in liquid, using Multimode VIII AFM equipped with a type E scanner (Veeco Instruments, Santa Barbara, USA) and silicon-tipped nitride cantilevers (Bruker, Camarillo, USA). Freshly cleaved mica was incubated with SEC-purified DNA pore solution (100 μL) for 15 min. Excess liquid was wicked off and replaced with 1×TAE buffer (100 μL) supplemented with 14 mM MgCl₂ and 4 mM NiCl₂.

Nanopore Current Recordings

For planar lipid bilayer electrophysiological current measurements, an integrated chip-based, parallel bilayer recording setup (Orbit 16, Nanion Technologies, Munich, Germany) with multielectrode-cavity-array (MECA) chips (IONERA, Freiburg, Germany) was used (Burns J. R., Seifert A., Fertig N., Howorka S. A. Nat. Nanotechnol. 11, 152-156 (2016); Del Rio Martinez J. M., Zaitseva E., Petersen S., Baaken G., Behrends J. C. Small 11, 119-125 (2015)). Bilayers were formed by spreading via a magnetic stirring bar DPhPC dissolved in octane (10 mg mL⁻¹). The electrolyte solution was 1 M KCl and 10 mM HEPES, pH 8.0. For pore insertion, a 2:1 mixture of cholesterol-anchored DNA nanopores and 0.5% OPOE (n-octyloligooxyethylene, in 1 M KCl, 10 mM HEPES, pH 8.0) was added to the cis side of the bilayer. A positive voltage of +30 mV was applied to facilitate pore insertion. Successful incorporation was observed by detecting the current steps. The current traces were Bessel-filtered at 2.873 kHz and acquired at 10 kHz with an EPC-10 patch-clamp amplifier (HEKA Elektronik, Lambrecht/Pfalz, Germany) with the PATCHMASTER software (HEKA Elektronik). Single-channel analysis was performed using Clampfit (Molecular Devices, Sunnyvale, Calif., USA).

Release Assays with Fluorophore-Filled Vesicles

Giant unilamellar vesicles (GUVs) were formed by adding a solution of DOPE (0.3 mmol, 50 μL) and DOPC (0.7 mmol, 550 μL) to an oven-dried round bottom flask (10 mL), removing the solvent via vacuum using a rotary evaporator for 20 min, and drying the thin film under ultra-high vacuum for 3 h. A solution of sorbitol (1 M, 1 mL) containing PEG³⁵⁰-FAM (10 μM) was added to the flask, and the solution was sonicated for 30 s to form fluorophore-filled vesicles. After 20 min incubation, a portion of the GUV suspension (1 μL) was added to PBS (200 μL) within an 8-well glass chamber (LabTek). After settling the vesicles for 5 min, a mixture of NP/OPOE (12.5 μl 0.5% OPOE, 37.5 μl PBS, 100 μL SEC-purified NP, incubation at 35° C. for 30 min) or OPOE/anchor-cholesterol strand (12.5 μL 0.5% OPOE, 37.5 μL PBS, 100 μL SEC-purified anchor-cholesterol strand, incubation at 35° C. for 30 min) was added. The fluorescence images of GUVs were collected using excitation at 515 nm and the appropriate emission filters.

Example 1: Assembly and Characterization of Wide Channel DNA Nanopores According to the Present Invention

An embodiment of a membrane-spanning DNA nanopore 10 according to the present invention is shown in FIG. 1 . In this embodiment, DNA duplexes are squarely arranged and interlinked to create a wide channel lumen having interior dimensions of approximately 7.5 nm×7.5 nm. As outlined in the “Nanopore current recordings” section above, the wide pore successfully inserts into lipid bilayers and shows considerably less closures at high voltages than prior art smaller diameter DNA nanopores (see, for example, Douglas S. M., Marblestone A. H., Teerapittayanon S., Vazquez A., Church G. M., Shih W. M. Nucleic Acids Res. 37, 5001-5006 (2009); Zheng J., et al. Nature 461, 74-77 (2009); Rothemund P. W. Nature 440, 297-302 (2006); Fu J., et al. Nat. Nanotechnol. 9, 531-536 (2014); Burns J. R., et al. Angew. Chem. Int. Ed. 52, 12069-12072 (2013); and Seifert A., Göpfrich K., Burns J. R., Fertig N., Keyser U. F., Howorka S. ACS Nano 9, 1117-1126 (2015)).

The nanopore 10 was designed with the CaDNAno software (Burns J. R., et al. Angew. Chem. Int. Ed. 52, 12069-12072 (2013)) and when assembled comprises squarely arranged DNA duplexes 12, 14. The DNA duplexes 12, 14 are interlinked by cross-overs 16. In this embodiment, the pore 10 has a total height of 46 nm and features and has a square design in cross-sectional profile parallel with the membrane surface. The external side length of the pore 10 is 22.5 nm.

In this embodiment, the cap region 18 is 35 nm in height and comprises a generally-square channel or lumen 19 surrounded by an upstanding pore wall 22. The pore wall 22 is composed of multiple duplex layers to increase structural stability (FIG. 1 ). In this embodiment the pore wall 22 is composed of three duplex layers, although depending on the requirements of the pore the pore wall 22 may be one; two; three; four; or five or more duplex layers thick.

In the membrane-spanning region 20, the pore wall 22 may comprise fewer duplex layers than that in the cap region 18 to decrease the overall pore size and hence lower the energetic costs of membrane insertion. In the embodiment shown in FIG. 1 , the pore wall in the membrane-spanning region is two duplex layers thick.

The membrane-spanning region 20 carries lipid anchors 24. The lipid anchors 24 are composed of cholesterol.

As shown in a cross-section (FIG. 1C), the pore lumen 19 is 7.5 nm×7.5 nm and features at its top a wider opening to help facilitate entrance of molecules. When inserted into a membrane (FIG. 1A), the pore 10 is expected to enable transport of protein cargo 26 from the cis to the trans side of the membrane.

The pores were successfully assembled via the DNA origami ‘scaffold-and-staple’ approach (Rothemund, P. W., Nature 440, 297-302 (2006). In this important route to DNA nanostructures, staple polynucleotides direct the folding path of a long single stranded DNA scaffold. The 2D-DNA map and DNA sequences of exemplary component strands are shown in FIG. 2 and provided in Table 1, respectively.

The sequence of the DNA scaffold is from M13mp18 DNA (SEQ ID NO. 1)

TABLE 1 Sequences of DNA staple, adaptor, and cholesterol-modified anchor polynucleotides. All DNA sequences are from the 5′ to the 3′ terminus. The names of staple and adaptor polynucleotides indicate their position within the 2D DNA map of the pore (FIG. 2) described as helix and base pair coordinate where the strands′ 3′ terminus is positioned within the numbered DNA helix of the nanostructure. The name of two cholesterol-modified anchor strands indicates whether the cholesterol-TEG modification is attached to the 5′ or 3′ terminus. Adaptor strands have two sequence parts which hybridize either to the DNA staple strands or the cholesterol-modified anchor polynucleotides (highlighted in bold). Adaptors that carry the latter sequence part at the 5′-end hybridize to 3′-cholesterol anchor polynucleotides. Conversely, adaptors with the sequence part at the 3′-terminus hybridize to 5′-cholesterol anchor polynucleotides. Adapter strand 21[64] features both a 5′- and a 3′-terminal  hybridization segment. Name: SEQ ID position] Sequence No. Helix [base Staple strands 2 47[136] GGAACAAATCATATATCCCACAAGCTTACCGAAGCTTGATTTCGGTCG 3 71[156] AAATGTTACAAAATCGCGCAAAA 4 35[125] TAACACCACCAGAGCCACC 5 44[190] GAAGGGTTAGAACCTTATACTTCTGAATAA 6 45[115] CATTAAGTAAGCATGAGCGCTCCCTGAACTCTGG 7  0[102] CCAAGCGTTGAGCCAAGGTGAATGTCA 8 33[109] GTGCATTAATTAGCTCGAATTCGTAATC 9 52[111] AAAATTCGAACCAATACTCCCGACAAAGCACTC 10 62[135] ACGGCCAGTAGCTGTTTCCTGTGTGAAATTGTTAT 11 65[149] CTCAAGTGTAAGAATCATAACCGAGTAAAAGAACG 12 22[90] TAAGAAGAAAATCTACAAAGCCGGAGACAGTCAAATCAC 13  4[94] GGCTCCATTAATTGTCGAAC 14 34[164] TATTTCAGAGCGGATGGTTGCTTTGACGAACGCT 15 32[122] GGCTGAATTTAGCCTTGAGT 16 44[106] AAAGTTACCAGATAAAAGAGGACTAAAGCGATTATA 17 48[156] TGAGTCAGAAGGAGCGGAATTATCATCATATATAATCAGC 18 57[131] CGCACTTCCAAGAAGATAAGTGTATAGCCGGATTAGGA 19 31[125] AACAACATGTTCAAGAGAACGGAATAGG 20 25[133] GATAATTTGCCTTGCTAAAGCGAATAAT 21  0[82] AACTAGCAACGGCTACAGAGTC 22 10[118] AACAGTGAGACTCCTCAGCTAATGCAGATAAGGCT 23 56[166] AGAACCCCCAGTCACACCAATCAATCCT 24 70[130] AGAACGTGGACATCAAGTTTTCAATTATTGCTCCTGCT 25 13[128] ACCGGAACCAGTAGCGTAATT 26 31[112] ATTTAACAAGCTGGCGAACTGTTGGGAAGGGCCGG 27 20[94] AGAACTGATAAAGCTAAAGGGTGAGA 28  2[170] AACCATCGCCCCACTACGACTTATTACACCAGCGC 29 43[141] TTTATTTCGCAATCAATAGGAGGGAGGGCACC 30 27[113] ATCGAGAAGATGGGCGGAACAAACGGCGGATTGAC 31 10[83] TATTTCGGATAATCTTGACAATTATAT 32 62[114] TGCGGGTTTTCCCAGTCACATTACGCCACGCC 33 12[167] CCACCCTCAAGACAAAGAGTC 34 24[122] CAAATAAGAAAATCGTAAAAAACAAGAGAATC 35 25[157] CTAAAGCATCACAATATCTGGTCAGTTGCACTAACGCAGCCTTTACAG 36 19[141] GAACGTAGAAATTAAACGGGAATACACT 37 52[193] CCACGCTGAGAAAGGAATTG 38 28[175] ATGTAGAAACGACCAGAATACCTACATTTTGATTA 39  1[120] CATGAGGATGCAGGGACGAGG 40 66[132] AACGCGCGGGGAGGCAACAGCCTACCTTTGTCGC 41 26[114] AACGGAACGCCGCCAGCTTT 42 56[185] CATTCTGGCCTCTTTAAT 43 20[111] GATAGCCGATCAGCTTCCGCTT 44 35[157] CGCAGAGCCACGCCACCCTCAGAACCGACCAA 45 13[141] CCCTCATAGGTAAATGCTGAACAATCGGCC 46 52[143] AACCTCAACCACCAGCGTATTCTATATTTTCACCT 47 32[90] TGCCTCATTAAATGCCCCCTGCC 48 19[125] AAATTAAGAAATGACCCTGTAATA 49 41[115] GTAGCGGAAATTAACGGAATAGACCCCCAGACTTTTT 50 64[164] AGCCGGAAGCATAGTTAGAATTAGTTAATCCAAT 51 57[108] GACAGCCGGAAACCAGGCAAAGCGC 52 39[125] GCAATATTGACTCAACATGTTTTAAATATGCA 53 53[164] GGCGGTCAGTATTAACACCGGCGCGAA 54 31[88] GAATCGTCGACTGGATAAACATGATAGTAC 55  4[146] CTAGTCAAAAACGTCTTTCCCCTCAATCCTTGCTG 56  7[57] ATGAACGGTGTACAGACTTTGA 57 37[133] AATTCAATATAAAATCGG 58  1[155] AATGCACGCATAAAGAACTGGAAATAGCATATTTCAAAAA 59 22[122] ACAAATATCAGTAATGCCGATTCAACCGTTCTAGC 60 30[138] CGACGGCTGGTAATATCCAGAACAATACGCTCAATAAT 61 54[110] CATCAACTCTCCGTGGCATCGTAACCGTGCATCTGC 62 14[162] TGATAGCGATAAATTACCTTAGCCCGAAGTGTTGTTCCAGTTTCACT 63 46[120] TGAGTAATGAATATGATGAGAGGGTA 64  2[135] CTGAGGCTAGTTTCCAATACATACTTGTCACAAAT 65  9[104] AGAGGCTGCCCGTATATCAGCCAT 66 45[133] CTTTTGCGGGAAATAAAGATAACG 67  8[89] CGCCACCCTCAGAACAGGCGCATAGGTTCATCAAGAGACCTATTATT 68 53[96] CATTTTTTCATTAAATAAAGGAATGAGAT 69  4[114] TTGCAATCCAAAATAAACAGAAGATTGATTTTGTT 70 54[167] CTAAAACACGCGCCCAAATCAGAT 71 42[190] GAGACCCAATTCTGCAGTACCTTTTACATC 72 61[91] CGATTAAGTTGGGAATCCCCCTGACCATAAATCA 73 47[108] CATCTGTAGGTAAGGGTAAT 74  0[170] CACCAACCTAAGAAACGTC 75 32[170] ATGAGCAATACAGTGTTTTTATAATCACACAATTCACG 76 50[132] CGGTACGATTTTTGAGAATTATCTTAAACAGCCC 77 12[110] CCAGCATTGGAAAGCCGTAAAATCAGGTCTTGCCCGCTTGGGCGCCAGGC 78 38[178] AATTTCATTTGGCTTAGATGAAA 79 37[109] TTCCGGTCCACTTCACCAGT 80 10[164] CAGGAGTGTACTGGTAATAAGGGTTTTGCTCTGT 81  8[107] ACTCTAGACGCGCCTGTTTACTTCTGGTGTATCGG 82 15[141] ATTTACATAAATTTCCCTTCAACCGCCTGGC 83 68[171] GCGACCACACCCGCCGCGCTCTACAGGG 84 24[106] ATTTTGCGGGACTCATAGTCCACCACCCCGT 85 37[89] AAATATCTAGCGCGTTCACCGACCGAAA 86 29[155] ATCCCATAATCGGCCGTAACAATAGAAGGCCCAG 87 16[170] AGACAAAAGGGAAAATTAAAATAC 88 64[133] GCCTAATGAGTGAGGTCGTGCCGGTTTGAATTATA 89 24[98] TTAGACGTTAGCAAAAAAAA 90 60[167] GAGTAGAAACCGTTGTCGTTATACAAAAAGCCATA 91 13[109] CACCGGCAAAAAGATTAAG 92 17[125] ATTATAAAGGTACATCCAATAAATTGCGTAGA 93 15[69] TGCTCATTCAGTGAATAAGTTTCATCGGCATTTTCGGTCACAACG 94 48[193] TAAATCCTTTTTATCAGA 95  6[129] AGCATTCCACATGGGATTAGTTA 96 18[148] GCAAATACCCAAACCGATATAACCGATAGAACAA 97 67[113] GAGACGGAGGCGGTTAGGTTGGGATACCGACGCAG 98 48[110] GCTATTTCCTGAGAGAAAGTCAGATTTA 99 49[139] TAATATTAGACGGTGTTTAACAAGGAATTCAACTTTC 100 12[82] CAGACGATTGGCCTTGCATTA 101  0[134] AAAACACCACCAGTAGAAGGTAAAAAGAAGATTATTCAT 102  6[94] GATCTAATCTTTCCAGGAATACCGAAAGATTCCGG 103 59[91] CATTCGCCATTCAAATGTTTAATAAATATAACAGTTAAGCCAGAATGAC 104 50[183] TAAAATATCTTTAGGAGGCAAATCAACAGTTG 105 71[87] GGGCGATGGTTTTGCGGATG 106  4[162] GAATAGAAAGGTTGCGCCGAAAACAGGTAAGCCCAGTT 107 18[90] TGGTGGCATCATAAAGCCTCAG 108 40[190] AACGGATTCGCCTGAAACAGTTGATTAACA 109 37[149] CCTTATCAAAATCAGAGCCCACCCTCAG 110 69[135] CAAAATCCCTTATAAATGGCGCTGGAGAAT 111 52[183] AGCCAGCAGCAAATGAAAAATCTACAATTTTATACCAACG 112 43[157] CAGTTGCACGTAAAACAGAGAAGCCTTATAGC 113 50[143] CCGTCAATATATCAAACAGAGCCTTAGTTGCTACT 114 29[165] AATTTACGAGCAGTACCGAAGCCATTGACTTGCCT 115 31[157] CAGAAAGTAATTCAGTACCAGGCG 116 59[108] GCGCAAAGGGGGATGTGCTGCAAGG 117 13[62] CAAATTAGCCCCCTTATTAGCGTTTGCAGGT 118 55[150] AGACAATATTTTTGAATG 119 39[100] GGATGAGGTCATCCCACTACGTGA 120 20[130] TTTCGCAATAAAATCATACAGGCAAGGCAAAGAATAAA 121 50[91] TTGAGGCTATCAGGTCATTGTTGAGAGAATCTA 122 29[88] TTGCCAGACGAGAGGCACCGCCACATAGTAAGTAA 123 68[158] GCGCTAGCAAAAGAATTTTTAATAAGAAAACCGAC 124 26[100] GAGGCCCTCAGAGTAGCGTAAC 125 41[147] GTACGAACCGATTAAAATTCAAGGCAAAAGTAAAATACGT 126 46[156] ATTTTTAGAACCCGAAACCACAACATTATCATTTTGCAGA 127 23[157] ATAAACTAATAGATTTAGAAGTATTAGTTTTAAAAATAATAAG 128 30[146] CCAAGAAAAATCGTCTGAAATGGAAGCCAGCTTTCGAT 129 33[131] AGAATAATTTAATAGCTGCATTAATGGTGCTTTC 130 19[93] CTTATGCGAAATTAAGCAAATTCTACTA 131 48[175] GTTATTAAACTTTACAAACAATTC 132 34[138] CTAAAACACCGAGCCTGGGGT 133 36[178] GACGCTGAGAAGAACGCGACGCGT 134 29[101] TAAAGGAGGTTAAGTATTA 135 49[133] GATTGATAAATAGAGATAATTTAAATGCAATGCC 136 70[170] TTGGATAGCCGGCGAACGTGAAGGGAA 137 42[132] GGTCAATAACCTGTTTAGCTATAGCATTAGGCAA 138 67[157] GCGAGTGAATTTGAAAACAAACCATCGCAAGG 139 11[116] TCTCTTAATTGAGAATCTTGTAACGCCACTGCAGGTCGACTCTAGAGGA 140 23[109] TCCAAAATCTCTAAATGAA 141 12[99] AGGAAAGCGGATTGCATCTTTGCGTATTTCCAGTC 142  3[64] TTGTGTCGAAATCCGCAAAGACAGCATATAAAGCCTGCGG 143 36[162] AATCGTAACCAAAGGAGCGG 144 58[124] CACCGTCAACAATACGGGTATCAGGGATAAGGCG 145 63[121] ATGGTCATGCCAAGCGGCGTTAAAGTAG 146  2[92] ACCCTCAGCAGCGGACAGCCTAAAGG 147 71[108] ACCATCACCCAATCCAACGTACCTTTAAAATAA 148 67[96] TTTTTCTTGCTGGTTTGCC 149 43[105] ATAGTAGTATTTTCAAAGACACCTTCATTAATTTG 150  8[164] GATAAGTGCCGTCGAGAGGGTCCCATGTACTGTC 151 57[91] CAGTTTGAGGGGACAAAATAGGGGGG 152 16[155] ATTCGTGTCTGGTTTGACCATTAGATACTCAGGTTTTTA 153 44[167] AAAATTATATGAATATACAGTAACGAACGAGTCAA 154 46[190] GATGGCAATTCATCAATTCCTGAGCCCGAAC 155  1[72] AACGAGGGGGAGATTTGTATCATCAAAGC 156 18[114] CATAAGGAAACGTTAAAGGGCTTTCGATCACG 157 42[167] AGATTTAGAAGTTTCATTCC 158 11[133] CCAACCAGAACCTATATGTCTGAGAGATGATTGCC 159 40[162] CAACGGAACCCTAAAGGGAGCCCCCGAGACGGGGA 160 54[122] GTAATCAAAAATAATTCGCAATTGTAAATCAA 161  0[90] CAACGAGTAGTAAATTGCATTTGGGGCAATTGCTG 162  8[118] TGTATCACTCATTTTTAAACCAAGTACCAACCGAC 163 50[193] GAAGGTTATCGACAACTCGT 164  6[140] ACAACAATAGGAATGATATAACCTGAACAGACGA 165 68[131] CCTGAGAGAGTTTGGTTCCGTGTGAGTGCCTGA 166 15[108] GGAATGCAGCTTCAAAGCGAGCAGGCGAGAAAAACCGTCTATCA 167  4[126] AATTTTTGGTGAATTACTGA 168  5[112] TTTTCTGTAGACAGCCGGTTTTGATAGCG 169 15[157] CCGAACGAAAGTATGGTTTGCAGTATGAACGT 170 55[132] CGTAGTCTGGCCGCCGTTTTAGAACGCGGCAAGCCCGCCTGT 171 63[155] CCGCTGTGAGGCCATTACTAGAAATTCTTGCTTTTGATGATA 172 41[172] ATATTTGCTTTGTTACATTTAAGGG 173 21[101] CGTAAGATTCAAAATCGGTTGTACCAATAGCA 174 71[128] TTTTGGGGTCGAGGTGCCGTAAAGGGAACAAGCAAACATCGGAAA 175  5[144] AACAGTTTCAGTACAAATTTTGCACTTATCCGAGA 176 57[119] CCTCAGGAAGTTGGTGTACAAGCAATTCCT 177 59[163] CCGCCCAAAAGGTTAATAAGAGAATATAAATCAT 178 20[146] TATAATTGAGTGAAGCGCACATTTGAGGATTAGAG 179 46[167] CTGGGATACGCAAATTGTTTGGATACCATATCAGA 180 60[185] TAATAACATCCAACAGGAAA 181 56[154] ACCATCATTACTCGCCATTAAACAGAGGTGA 182 22[172] AGAGAATAACATAACAATGACAAACAATGCATGATTA 183 61[140] TCTGTCCATCACGCAAATTAGAACTCAACGAGC 184 37[125] TATACAGAATCAGCAAAATTCATCTTTAGTTT 185 41[104] AATATAATTTTGATAATAGAGAGTCAAAGGGCAAATCCTGT 186 62[185] AATCCTGAGATTCTTTGATT 187 34[148] ATCTTCATACATGACCAGTATGGCATTTTACTATC 188 67[136] CTTCAGTGTAGCGGTCGCACGTATATGCAAATTTC 189 34[178] AACTTTTTCAATGTTTAGTACAAACATCACACGGAACGGTACGCC 190 60[145] GGCCTTGGCCTCTTCGCTGACGTTGTCGCTCAACATA 191 66[170] ACTGAGCTAAACAGGAGGCGATTTTAG 192 53[147] AGATAAAAAATACCGAACGAAATGATACGTGGCAC 193 63[91] TCCCCGGGTACCGGCGTTGCGCTC 194 38[114] CCTTTAGAGCCAAGTTTGCCTTTAGCGAAAAT 195 28[146] TTTTCGTAGGATGAAAGCGTAAGAGGATAGGTCACGAT 196 27[93] CAGTGAGCGAGAATCAGCT 197 28[140] TATCATCCTTATTTACATTGGCAGATTCATTCTG 198 54[176] CTGATAGCCGCTATTAGAACAGAGAT 199 65[111] GGGAAACCTCTAACTCAATAAATAATTGCA 200 50[98] ACCCATCAGTTTAATAAAAATCGGTTTAACATTTTA 201  6[162] AGTTTCGTCACCAGCGGAGCTAACGAGTGAAA 202 13[96] TCATAATCTCAGACTGGCGTTTTAATTCG 203 26[122] TTTAGCCTTAAACGTTAATTATAAGCAAATATTTAGAA 204 41[141] ACTGAGGCGAATGATGAAAAGTCCACTATTAA 205  6[64] AAGAGGACAGGCGCAGACGGTCAATTACTTAGCCGGAACGA 206 12[142] AGCCGCCGTAAGCGTCTGAC 207 55[88] TAACAACCCGTCGGATATTAAATGACGACGATTAC 208  5[80] AACTGTCGAGTTTCGACAG 209 49[117] AGCACTAGCATGTCAATCAAAAAACAGGCCAT 210 58[185] CGCTCATGGATAATAAAAGG 211 59[131] CGGTGCAATAAACATGTAATTGGGTCAGTCCGTT 212 16[90] CCGTAGAGCTTGCGAGCTGAAA 213  9[137] TTAGCGGTTTTAACGTAGGCAGAAAAGCCAAAAAG 214 69[113] TTGATGGGCAGCAAGTGTAAATCTTAAC 215 38[146] CAGACCATTAGATAGCAGCACCGTAATCGCCT 216 25[91] CGCCATTTTGTTATAATCAGAAAAGCCCCTATGT 217  2[105] TTGCGGGATCGGCTTTGAAACGGGCTTGAGA 218 36[114] CTCCGGAACCAGAGCCGCCG Adaptor strands 219 33[64] CTCCGTCTATCTTTTTGTTCAGAAAACGAGAATCAAA 220 39[64] CTCCGTCTATCTTTTTTTTCAAACTCCAACAGTATCA 221 25[64] CTCCGTCTATCTTATGACTGACCAACCCGCCA 222 27[64] CTCCGTCTATCTTCATAACCCTCGTTTAC 223 31[64] CTCCGTCTATCTTTTTATACTGCG 224 17[64] CTCCGTCTATCAGAAACACCAGAAAGTAC 225 19[64] CTCCGTCTATCTTAATCATTGTGAATTAC 226 29[64] CTCCGTCTATCTTTTTAAGAAGTT 227 37[64] CTCCGTCTATCTTTTTTTTGACTTC 228 35[64] CTCCGTCTATCTTTTTTTTATAGTCAGAAGCAGGTTGAGGCCATCTTT 229 23[64] CTCCGTCTATCTTTATCTCCATGTCATAAGGG 230 21[64] CTCCGTCTATCGTCAGGACGTTGGCGGAACAACATTTGCTACGTCAGC 231 65[93] ACTTACCCTGACTATTTTTTTTTTGCTACGTCAGC 232  4[82] CTGTACAGGTAACATTCAACTATTGCTACGTCAGC 233 43[85] AGGTTTAATTTCAACTGCTACGTCAGC 234 45[85] AGCGCTCATTATACCAGCTACGTCAGC 235 41[85] GCTGCTTGCCCTGACGGCTACGTCAGC 236  9[85] CTGAGCGTCCATTTTTGCTACGTCAGC 237  7[80] CCCTCAGATTTTGCAATTTTTGCTACGTCAGC 238 11[71] TCACAAACAAATAAATCTTTAAACATTTTTGCTACGTCAGC 239 25[83] ATACAAGCAACACTATTTGCTACGTCAGC 240 67[93] TGGAGGAAGCCCGAAATTTTTTTTGCTACGTCAGC 241 69[93] CCAACCAGACCGGAAGTTTTTTTTGCTACGTCAGC Anchor strands 242 Cholesterol 3′ GATAGACGGAG-TEG-Chol 243 Cholesterol 5′ Chol-TEG-GCTGACGTAGC

Lipid anchor-free variants of the nanopore (NP^(−c)) may be assembled first and then converted into a lipid-modified pore (NP) by hybridizing to sticky ends within the transmembrane region polynucleotides carrying the cholesterol anchors. Assembly of NP^(−c) followed an optimized annealing protocol (see “Assembly” section below). The fabrication outcome was analyzed by gel electrophoresis to yield a single defined band as shown for cholesterol-free NP^(−c) (FIG. 3A, panel −SDS), implying a homogenous population of folded products. The pore band migrated at a different height than the scaffold strand (ss) (FIG. 3A) suggesting complete assembly. Pore NP with cholesterol anchors also led to a defined band when analyzed in detergent SDS (FIG. 3A, panel +SDS) to suppress the known streaking likely caused by hydrophobic interactions with the gel matrix (FIG. 3A, panel −SDS) (Langecker M., et al. Science 338, 932-936 (2012); Burns J. R., Al-Juffali N., Janes S. M., Howorka S. Angew. Chem. Int. Ed. 53, 12466-12470 (2014)).

DNA pores with a molar mass of 4.87 MD were purified via size exclusion chromatography (SEC) (FIG. 4 ) from excess staple polynucleotides and used for the biophysical characterization.

Atomic force microscopy (AFM) was applied to determine the dimensions of the synthesized DNA nanopores. The samples were incubated with Mg²⁺ and Ni²⁺ to adsorb the negatively charged nanostructures onto negatively polarized and atomically flat mica substrates. AFM micrographs of pore NP^(−c) featured chain-like structures composed of multiple pores (FIG. 3B, top panel; FIG. 5 ). Related higher-order assemblies have been observed for other surface-adsorbed DNA nanostructures that are held together by counter-ions or blunt-end stacking (Burns J. R., Seifert A., Fertig N., Howorka S. Nat. Nanotechnol. 11, 152-156 (2016); Aghebat Rafat A., Pirzer T., Scheible M. B., Kostina A., Simmel F. C. Angew. Chem. Int. Ed. 53, 7665-7668 (2014)). It is certain that the multi-component chains solely formed due to substrate effects since SEC (FIG. 4 ) and gel electrophoresis (FIG. 3A) confirmed the monomeric nature of pores in solution. As shown by AFM analysis, the DNA nanopores of high aspect ratio adsorbed in lying orientation onto mica to maximize cation-mediated interaction with the negative substrate surface. Indeed, AFM profiles of separate DNA pores (FIG. 3B, middle and bottom panels) yielded an average elevation of 10.2±1.1 nm (n=19) in agreement with a lying pore that is compressed by the AFM cantilever tip (Burns J. R., Seifert A., Fertig N., Howorka S. A Nat. Nanotechnol. 11, 152-156 (2016)). The value of 10.2 nm is within the range obtained for other DNA structures also six duplexes thick (Schmied J. J., et al. Nat. Protoc. 9, 1367-1391 (2014)). Compression of the pore accounted for its square appearance (FIG. 3B) since DNA duplexes flanking the pore lumen are squeezed out. The squares had a side length of 36.0±10.5 nm (FWHM, n=19, FIG. 3B, bottom panel) which comprises both the broadened nominal width of 22.5 nm and the height of 35 nm of the pore's cap region. The majority of the transmembrane region of NP^(−c) without cholesterol-containing strands is only 2 duplexes thick (FIGS. 6C and D) and hence less clearly imaged in AFM.

The anchoring of cholesterol-tagged NP into lipid bilayers was established using a gel shift assay. In line with anchoring, the band for NP was upshifted and co-migrated with small unilamellar vesicles (SUVs) that were unable to enter the gel to remain in the loading slots (FIG. 8C). Increasing amounts of SUVs led to a complete conversion to the upshifted DNA band (FIG. 8C) implying that all pores interacted with the bilayer. By contrast, NP^(−c) without anchors did not produce any gel shift (FIG. 8C), in agreement with the need of cholesterol for membrane insertion. Pores with half the number of cholesterol anchors led to an incomplete gel shift (FIG. 7 ).

The membrane-spanning nature of nanopore NP was confirmed with single-channel current recordings. Individual pores were inserted into a planar lipid bilayer (Del Rio Martinez J. M., Zaitseva E., Petersen S., Baaken G., Behrends J. C. Small 11, 119-125 (2015); Goyal P., et al. Nature 516, 250-253 (2014)) and a potential was applied across the membrane to induce flow of electrolyte ions (Burns J. R., Seifert A., Fertig N., Howorka S. Nat. Nanotechnol. 11, 152-156 (2016)). Under standard electrolyte conditions, a constant current of 49.5 pA was observed (FIG. 3A) at a potential of +20 mV relative to the cis side of the pore. The corresponding conductance distribution of 32 pores had a maximum of 2.29±0.26 nS (FIG. 3B). In agreement with the wide pore lumen, the conductance is 2.6-fold higher than the reference DNA pore of 2 nm diameter (Langecker M., et al. Synthetic lipid membrane channels formed by designed DNA nanostructures. Science 338, 932-936 (2012)). Calculating the theoretical conductance based on the geometry of NP yielded a value of 6.7 nS. But this is too high because the basic assumption of the model—a constant mobility of electrolyte ions as in bulk solution—is not valid for ionic transport in nanoscale confined pores with negatively charged walls (Ho C., Qiao R., Heng J. B., Chatterjee A., Timp R. J. Electrolytic transport through a synthetic nanometer-diameter pore. Proc. Natl. Acad. Sci. USA 102, 10455-10450 (2005)). Given the lower experimental conductance, it seems unlikely that ionic leakiness across the 3 duplex-thick pore wall plays a major role. Leakiness has previously been found in simulations for a single duplex pore wall (Maingi V., Lelimousin M., Howorka S., Sansom M. S. ACS Nano 9, 11209-11217 (2015)) and for ionic transport across a layered duplex structure that was oriented perpendicular to the electric field (Yoo J., Aksimentiev A. Proc. Natl. Acad. Sci. USA 110, 20099-20104 (2013); Plesa C., et al. ACS Nano 8, 35-43 (2014)) rather than parallel as in the case of NP. Voltage ramps established that NP was of ohmic behavior (FIG. 3C) as expected for a symmetrical pore. Above 30-40 mV, the pore could switch to a lower conductance state. The magnitude of the conductance drop at 30% (FIG. 3D) is considerably smaller than the value of 80% for previous DNA nanopores (Seifert A., Göpfrich K., Burns J. R., Fertig N., Keyser U. F., Howorka S. ACS Nano 9, 1117-1126 (2015); Burns J. R., Seifert A., Fertig N., Howorka S. A biomimetic Nat. Nanotechnol. 11, 152-156 (2016)). Furthermore, the probability of switching the conductance state is much lower for the new pores. Quantified as probability for the higher conductance state, the new large pores were open 100% at 20 mV and 75% at 100 mV (FIG. 3E, two left bars, “native”) compared to 80% and 20%, respectively, for small DNA nanopores. The native and open state of large DNA pores could, however, be perturbed by repeated voltage switching to achieve a memory effect that results in a lower open probability (FIG. 3E, two right bars, “perturbed”).

Pore NP was able to channel molecular cargo across the membrane as shown by the release of a fluorescent probe from lipid vesicles. Giant unilamellar vesicles (GUVs) were filled with poly(ethylene glycol) PEG³⁵⁰ that was coupled to fluorescent dye fluoresceine amidite (FAM). Unlike fluorescence-tagged protein, FAM-PEG³⁵⁰ of 2.4 nm diameter (Merzlyak P. G., Yuldasheva L. N., Rodrigues C. G., Carneiro C. M. M., Krasilnikov O. V., Bezrukov S. M. Biophys. J. 77, 3023-3033 (1999); Krasilnikov O. V., Rodrigues C. G., Bezrukov S. M. Phys. Rev. Lett. 97, 018301 (018301-018304) (2006)) could be packed at high density into GUVs giving rise to a strong visual signal in fluorescence microscopic images (FIG. 10 , left panel); the size distribution of GUVs was within the norm (Moscho A., Orwar O., Chiu D. T., Modi B. P., Zare R. N. Proc. Natl. Acad. Sci. USA 93, 11443-11447 (1996)). Addition of cholesterol-tagged NP led to a stark decrease in fluorescence while leaving the SUV shape unaffected (FIG. 10 , right) strongly suggesting that the pore punctured holes into the membrane without rupturing it. In support of this interpretation, a negative control featuring cholesterol-modified DNA polynucleotides did not change the vesicle fluorescence (FIG. 104A, left) implying that membrane anchoring without pore activity does not lead to membrane puncturing.

The transport of protein along the lumen of the DNA channel (FIG. 11 ) was examined with single-channel current recordings. Trypsin with a molecular size of 4.3×3.8×2.3 nm (pI 10.1) was selected as test protein. Upon its addition to the cis side, two types of current blockades occurred that differed in their duration, τ_(off): Shorter type I and longer type II events (FIG. 5B). The blockades also had different levels of amplitude A (FIG. 11B). When each event was plotted with its τ_(off) and A as separate point in a scatter diagram, the two blockade types clustered into two different regions (FIG. 11C). Type I had τ_(off) of less than 1 ms while type II featured τ_(off) values between 2-200 ms with an average of 17.5±5.5 ms (N=3) obtained from the fit to the single exponential decay distributions. As other distinguishing characteristic, only type II had a defined amplitude A of 26.2±0.7% (N=3) when normalized to the open channel current I₀ (FIG. 11C) while A of type I ranged from 5 to 35%.

In line with related current signatures through inorganic pores (Wei R. S., Gatterdam V., Wieneke R., Tampe R., Rant U. Nat. Nanotechnol. 7, 257-263 (2012); Yusko E. C., et al. Nat. Nanotechnol. 6, 253-260 (2011); Oukhaled A., et al. ACS Nano 5, 3628-3638 (2011)) type I events are interpreted as short transient contacts of protein with the pore (FIG. 11A, bright green) while type II are seen to represent the movement of trypsin through the DNA channel (FIG. 11B, dark green). This interpretation is supported by (i) the agreement between the longer duration of type II events and the expected long transport process through the pore. A further support is (ii) the match between the experimental amplitude of type II events and the calculated blockade level of 30%. The latter value was derived from the ratio of the cross-sectional areas of trypsin and the pore lumen. Finally, the narrow distribution of the current blockade is in line with the homogenous nature of current blockades caused by proteins that migrate through a channel of largely constant width.

Transmission electron microscopy (TEM) images of the nanopores according to the present invention are shown in FIGS. 12 to 14 . FIG. 12 shows TEM images of the DNA nanopores of the present invention. FIGS. 13 and 14 show TEM images for representative DNA nanopore structures attached on SUVs. The consistency of the pore structure is clearly shown in the images.

Example 2: Preparation and Insertion of DNA Nanopores into Synthetic Polymer Membranes Synthesis of PMPC-b-PDPA Membrane Polymersomes

2-(Methacryloyloxy)ethyl phosphorylcholine monomer (MPC, 99.9% purity) (ex. Biocompatibles U.K.); 2-(diisopropylamino)ethyl methacrylate (DPA) and remaining chemicals were bought from Sigma Aldrich. Block copolymer PMPC-PDPA was synthesized by atom-transfer radical-polymerization (ATRP) following a published protocol. [L. Ruiz-Perez, J. Madsen, E. Themistou, J. Gaitzsch, L. Messager, S. P. Armes, G. Battaglia, Polym. Chem. 2015, 6, 2065-2068] Briefly, a solution in morpholinoethylbromoisobutyric acid ester (ME-Br, synthesis described previously) [H. Lomas, I. Canton, S. MacNeil, J. Du, S. P. Armes, A. J. Ryan, A. L. Lewis, G. Battaglia, Adv. Mater. 2007, 19, 4238-4243.] (0.190 g, 0.68 mmol, 1 eq.) in EtOH (5 mL) was placed in a round-bottom flask before addition of MPC (5.000 g, 1.70 mmol, 25 eq.). The mixture was stirred and further purged with nitrogen for 30 min and heated to 30° C. Then, a mixture of 2,2′-bipyridine (bpy) (0.223 g, 1.42 mmol, 2 eq.) and copper(I) bromide (Cu(I)Br) (0.097 g, 0.68 mmol, 1 eq.) was added under a constant nitrogen flow. The mixture was stirred for 60 min to yield a highly viscous brown substance and sampled with NMR to estimate the extent of conversion. Meanwhile, a solution of DPA (12.27 g, 57.6 mmol, 85 eq.) in EtOH (13 mL) was prepared and purged with nitrogen for 60 min in a separate flask. DPA solution was added to the polymerization mixture, and the reaction solution was purged for another 10 min and left overnight at 30° C. After 18 h, ¹H NMR analysis confirmed that the conversion was >99%. Upon diluting the reaction mixture in EtOH (30 mL), the solution gradually turned green, indicating oxidation of the copper-based catalyst. The mixture was passed through silica and the solvent was partially evaporated to give an opaque solution which was then dialyzed (MWCO 1,000 Da) against CH₂Cl₂, MeOH, and water (2 times each) for 8-14 h for each dialysis cycle. The polymer was freeze-dried and dried at 120° C. for 2 h under vacuum resulting in a yield of 13.3 g (77%). ¹H NMR analysis in mixture of CDCl₃/MeOD (3:1) determined the composition of the polymer to be PMPC₂₅-PDPA₇₂. Size-exclusion chromatography (SEC) established that the polydispersity index (PDI) had a value of 1.22. All polymersome dispersions were prepared by thin film hydration (J. Gaitzsch, D. Appelhans, L. Wang, G. Battaglia, B. Voit, Angew. Chem. Int. Ed. 2012, 51, 4448-4451). Typically, block copolymer PMPC₂₅-PDPA₇₂ was dissolved in a mixture of CHCl₃ and MeOH (2:1, 5 mL) and then dried under vacuum at 30° C. overnight. The resulting dried polymer film was hydrated in PBS (5 mL) and stirred vigorously. After 7 weeks under stirring, the polymeric vesicle suspension was purified through centrifugation cycles. Briefly, an aliquot (500 μL) was centrifuged for 10 min at 1000 RCF. The supernatant was further centrifuged for 20 min at 23 000 RCF, and the resulting pellet was re-suspended in PBS (450 μL) and sonicated for 20 min, yielding a monodisperse suspension of polymeric vesicles. The concentration of PMPC₂₅-PDPA₇₂ within polymersome suspension was measured by UV-vis spectroscopy. Typically, a polymersome suspension (20 μL) was diluted 1:10 in PBS, pH 2.0, and the absorbance was recorded at A=220 nm. The concentration of PMPC₂₅-PDPA₇₂ was calculated using a calibration curve of the polymer. The polymersome size and its distribution were determined by dynamic light scattering as described in section 1.8. To calculate the number of polymersomes in a unit volume of suspension, the concentration of PMPC₂₅-PDPA₇₂ and the polymersome size were used as input variables for a Matlab® script which is based on several equations. Following equation (1), the number of polymersomes, N_(p), is given by the experimentally determined total number of polymer chains, N_(c), divided by the number of polymer chains per polymersome, N_(agg).

$\begin{matrix} {N_{p} = \frac{N_{c}}{N_{agg}}} & (1) \end{matrix}$

Nagg is defined in equation (2) as the ratio between the volume of the polymersome hydrophobic membrane, V_(p), divided by the product of molecular volume of the PDPA block of the polymer, VPDPA, assuming a vesicle packing parameter p=1.

$\begin{matrix} {N_{agg} = \frac{V_{p}}{V_{PDPA}}} & (2) \end{matrix}$

The molecular volume, V_(PDPA), is defined as:

$\begin{matrix} {V_{PDPA} = \frac{M_{PDPA}}{\rho_{PDPA}N_{A}}} & (3) \end{matrix}$

where M_(PDPA) is the molar mass of the PDPA block, ρ_(PDPA) is the bulk density of the hydrophobic chain and is 1.02 g/ml, and NA is the Avogadro number. Insertion of DNA Nanopores into PMPC-b-PDPA Membrane

A solution of nanopores (0.005 μM, 25 μL, in 10 mM Tris, 10 mM NiCl₂ pH 7.4) was added to freshly cleaved mica and incubated for 5 min. at room temperature. The supernatant was then removed followed by the addition of buffer solution (25 μL, 10 mM Tris, pH 7.4) reducing the NiCl₂ concentration to ˜1 mM.

Encapsulation of Trypsin into Polymersomes, their Purification by Size Exclusion Chromatography and Characterization by UV-Vis Spectroscopy

Porcine trypsin (1000-2000 U/mg) may be encapsulated into polymersomes by electroporation following a published procedure (L. Wang, L. Chierico, D. Little, N. Patikarnmonthon, Z. Yang, M. Azzouz, J. Madsen, S. P. Armes, G. Battaglia, Angew. Chem. Int. Ed. 2012, 51, 11122-11125).

Briefly, a suspension of PMPC₂₅-PDPA₇₂ polymersomes (5 mg mL-1, 200 μL) and a solution of trypsin (25 mg mL-1, 200 μL) were gently mixed and placed into an electroporation cell. This mixture was subsequently transferred in the electroporator (Eppendorf 2510) and electroporated using 5 pulses of an AC electric field at a voltage of 2500 V with an interval of 30 s between each pulse. Polymersomes with encapsulated trypsin were purified from excess free trypsin by SEC using Sepharose 4B. The gel filtration medium (20% slurry in EtOH) was first washed multiple times with PBS and centrifuged (5000 RCF, 5 min). Sepharose was then packed into the chromatographic column and washed 5 times with PBS. The sample (400 μL) was applied to the column, and purified material was eluted by collecting 500 μL fractions. Isolated trypsin and polymersomes that was not exposed to trypsin were also purified to obtain the corresponding reference elution volumes (10-15 mL and 5-6 mL, respectively). The protein content in the SEC-purified polymersome fraction containing encapsulated trypsin was analyzed by UV-vis spectroscopy to obtain absorbance readings at 280 nm using a Carry Eclipse Varian spectrophotometer. To determine the PMPC₂₅-PDPA₇₂ polymer content, protein-free polymersomes that had been subjected to the same purification procedure as polymersomes with encapsulated trypsin were analyzed by UV-spectroscopy at 220 nm. The polymersome solution (20 μL) was diluted 10-fold in PBS at pH 2.0. The concentration of trypsin was PMPC₂₅-PDPA₇₂ was calculated from absorbance readings and calibration curves for PMPC₂₅-PDPA₇₂ polymer and trypsin in PBS at pH 2.0.

Enzymatic Assays of Polymersomes with Encapsulated Trypsin

Nanoreactor assays were carried out with SEC-purified polymersomes containing encapsulated trypsin. A suspension of polymersomes (14 μM trypsin, 0.62 mg mL-1 polymer, 50 μL) and solutions of DNA nanopores NP-3C or NP-0C (1 μM, 25 μL) and B-NAR-AMC peptide (1 mM, 25 μL) were added to PBS, pH 7.4 (100 μL). For the negative control, the DNA nanopore solution was replaced with PBS, pH 7.4 (25 μL). For the positive control without polymersomes and DNA nanopores, B-NAR-AMC peptide (1 mM, 25 μL) and trypsin solution (25 μM, 50 μL) were mixed with PBS, pH 7.4 (125 μL). To avoid immediate reaction in the positive control, a lower concentration of trypsin (500 nM, 50 μL) was used to lower the conversion rate. All measurements were recorded on a Carry Eclipse fluorescence spectrophotometer. The fluorescence emission of each mixture was monitored between 400 and 600 nm with λexc=380 nm.

In summary, described herein are rationally designed and structurally defined, unprecedented wide membrane pores. These pores provide a significant advantage compared to existing biogenic and synthetic channels and may have many potential applications including biosensing, synthetic biology, and DNA nanotechnology.

Unlike existing protein pores, the nanopores described herein meet the criteria of structural definition, a wide lumen of approximately 50 nm² cross-section, and a modular design for tunable size. The new DNA nanopore exceeds current DNA pores by a 10-times larger lumen and the higher structural stability, that is, a less extensive and less frequent lower conductance state. Since the lower conductance is likely caused by loose DNA termini that are normal in DNA nano-architectures, improved pores could be formed in future by ligating DNA termini to achieve a covalently closed structure. As a further advantage, the pore design is highly modular and takes advantage of tested structural principles in nanotechnology. It is realistic that the approach can be extended to build even wider or shorter DNA pores.

DNA pores that are wide enough to channel protein across a membrane open up many applications, for example use in sensors for protein detection. In other, equally exciting applications the pores could be adapted to create a molecular valve with a closable lid in order to regulate the flow of proteins across membranes (Burns J. R., Seifert A., Fertig N., Howorka S. A biomimetic Nat. Nanotechnol. 11, 152-156 (2016); Andersen E. S., et al. Nature 459, 73-76 (2009)). Valves may be used for drug delivery nanodevices (Mura S., Nicolas J., Couvreur P. Nat. Mater. 12, 991-1003 (2013)) that could be composed of stable vesicles with biocompatible polymer walls (Messager L., et al. Angew. Chem. Int. Ed. in press (2016); Howse J. R., Jones R. A., Battaglia G., Ducker R. E., Leggett G. J., Ryan A. J. Nat. Mater. 8, 507-511 (2009)). DNA nanopores may also help generate molecular machines that selectively transport cargo across the membrane (Franceschini L., Soskine M., Biesemans A., Maglia G. Nat. Commun. 4, 2415 (2013)). Finally, the new insight into how to anchor large negatively charged DNA channels into hydrophobic bilayers will strongly benefit the development of other membrane-tethered DNA nanostructures. Indeed, only recently have lipid-anchored DNA materials (Edwardson T. G., Carneiro K. M., McLaughlin C. K., Serpell C. J., Sleiman H. F. Nat. Chem. 5, 868-875 (2013)) been built to mimic functions of membrane proteins such as to control access of molecules to a cell's interior or to determine membrane morphology (Kocabey S., et al. Membrane-assisted growth of DNA origami nanostructure arrays. ACS Nano 9, 3530-3539 (2015); Czogalla A., et al. Angew. Chem. Int. Ed. 54, 6501-6505 (2015); Johnson-Buck A., Jiang S., Yan H., Walter N. G. ACS Nano 8, 5641-5649 (2014); Yang Y., et al. Nat. Chemistry 8, 476-483 (2016); Xu W., et al. J. Am. Chem. Soc. 138, 4439-4447 (2016); Perrault S. D., Shih W. M. ACS Nano 8, 5132-5140 (2014). These biomimetic structures are of interest in basic research, biotechnology and biomedicine (Howorka S. Nanotechnology. Changing of the guard. Science 352, 890-891 (2016); Chen Y. J., Groves B., Muscat R. A., Seelig G. Nat. Nanotechnol. 10, 748-760 (2015)).

Although particular embodiments of the invention have been disclosed herein in detail, this has been done by way of example and for the purposes of illustration only. The aforementioned embodiments are not intended to be limiting with respect to the scope of the appended claims, which follow. The choice of nucleic acid starting material is believed to be a routine matter for the person of skill in the art with knowledge of the presently described embodiments. It is contemplated by the inventors that various substitutions, alterations, and modifications may be made to the invention without departing from the spirit and scope of the invention as defined by the claims. 

1.-46. (canceled)
 47. A method for molecular sensing comprising: i) providing a semi-fluid or lipid membrane having a first side and a second side; ii) providing a nucleic acid membrane-spanning nanopore located in the membrane, wherein the nucleic acid membrane-spanning nanopore defines a central channel that allows for fluid communication between the first side of the membrane and the second side of the membrane, wherein the minimum internal width of the central channel of the nanopore is from about 5 nm to about 20 nm; and iii) measuring a change in electrical properties of the nucleic acid membrane-spanning nanopore in the presence of an analyte, wherein detection of a change in electrical properties is indicative of the presence of the analyte.
 48. The method of claim 47, wherein the semi-fluid or lipid membrane is selected from the group consisting of: a membrane comprising a lipid bilayer; and a membrane comprising a semi-fluid membrane formed of polymers.
 49. The method of claim 48, wherein the polymer forming the semi-fluid membrane is composed of amphiphilic synthetic block copolymers, suitably selected from hydrophilic copolymer blocks and hydrophobic copolymer blocks.
 50. The method of claim 47, wherein the change in electrical properties is a change in the flow of ions through the nucleic acid membrane-spanning nanopore.
 51. The method of claim 50, wherein the change in the flow of ions is from the first side of the membrane to the second side of the membrane.
 52. The method of claim 47, wherein the change in electrical properties is a change in electron flow across the nucleic acid membrane-spanning nanopore.
 53. The method of claim 52, wherein the change in electrical properties is measured across an aperture of the nucleic acid membrane-spanning nanopore, and the electrical property is selected from one or more of the group consisting of: tunnelling current; local potential; and capacitance.
 54. The method of claim 53, wherein the change in electron flow is measured by a field effect transistor (FET) device or a FET nanopore device.
 55. The method of claim 47, wherein the analyte is selected from one or more of the group consisting of: folded or unfolded proteins; DNA-protein constructs such as nucleosomes and polynucleotides such as deoxyribonucleic acid (DNA) or ribonucleic acid (RNA); polysaccharides; and synthetic polymers.
 56. The method of claim 47, wherein the analyte comprises a nucleic acid.
 57. The method of claim 56, wherein the nucleic acid is selected from one or more of the group consisting of: DNA; RNA; a synthetic nucleic acid; and a modified nucleic acid.
 58. The method of claim 57, wherein the nucleic acid is comprised of one or more nucleotides and the sequence of the one or more nucleotides may be determined from characteristic disruptions in the measured electrical properties over time.
 59. The method of claim 47, wherein the nucleic acid membrane-spanning nanopore comprises: a) at least one scaffold polynucleotide strand; b) a plurality of staple polynucleotide strands; and c) at least one hydrophobically-modified polynucleotide strand, wherein the at least one hydrophobically-modified polynucleotide strand comprises a polynucleotide strand and a hydrophobic moiety; wherein each of the plurality of staple polynucleotide strands hybridises to the at least one scaffold polynucleotide strand to form the three-dimensional structure of the nucleic acid membrane-spanning nanopore; and wherein the at least one hydrophobically-modified polynucleotide strand hybridises to a portion of the at least one scaffold polynucleotide strand.
 60. The method of claim 47, wherein the nucleic acid membrane-spanning nanopore comprises a membrane spanning region, wherein the membrane spanning region has a wall thickness of more than one DNA duplex.
 61. The method of claim 59, wherein the polynucleotide strand of the or each at least one scaffold strand, each of the plurality of staple polynucleotide strands, and the or each hydrophobically-modified polynucleotide strand comprises DNA.
 62. The method of claim 59, wherein the assembly of the nucleic acid membrane-spanning nanopore is via DNA origami techniques.
 63. The method of claim 59, wherein the nucleic acid membrane-spanning nanopore further comprises one or more adaptor polynucleotide strands, wherein the at least one hydrophobically-modified polynucleotide strand is hybridised to the nucleic acid membrane-spanning nanopore via the one or more adaptor polynucleotide strands, the one of more adaptor polynucleotide strands each having a first end and a second end, wherein the first end of the adaptor polynucleotide strand hybridises with the at least one scaffold polynucleotide strand, and the second end of the adaptor polynucleotide strand hybridises with the at least one hydrophobically-modified polynucleotide strand.
 64. The method of claim 63, wherein the polynucleotide in the adaptor polynucleotide strands comprises DNA.
 65. The method of claim 59, wherein the at least one hydrophobic moiety comprises a lipid, wherein the lipid is selected from the group consisting of: sterols; alkylated phenols; flavones; saturated and unsaturated fatty acids; and synthetic lipid molecules (including dodecyl-beta-D-glucoside).
 66. The method of claim 47, wherein the semi-fluid or lipid membrane is provided with a plurality of nucleic acid membrane-spanning nanopores. 